Regulation of the Complement System by Toxoplasma Gondii Surface Coat Proteins Srs25, Srs29c, and Srs57

REGULATION OF THE COMPLEMENT SYSTEM BY TOXOPLASMA GONDII SURFACE COAT PROTEINS SRS25, SRS29C, AND SRS57

A Dissertation submitted to the Faculty of the Graduate School of Arts and Sciences of Georgetown University in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Microbiology and Immunology

Patricia M. Sikorski, M.S.

Washington, DC August 19, 2019

REGULATION OF THE COMPLEMENT SYSTEM BY TOXOPLASMA GONDII SURFACE COAT PROTEINS SRS25, SRS29C, AND SRS57

Patricia M. Sikorski, M.S

Thesis Advisor: Michael E. Grigg, Ph.D.

ABSTRACT

Very little is known about how the protozoan parasite Toxoplasma gondii overcomes the complement system to establish infection. In this dissertation, I found significant differences in C3b deposition between T. gondii Type I and Type II strains.

Taking both forward and reverse genetic approaches, I identified three Toxoplasma surface proteins that regulate complement deposition: SRS29C and SRS57 were identified as C3b acceptors, whereas SRS25 limited the level of C3b on the parasite surface. Serum resistance was achieved by the recruitment of host regulator proteins C4b-binding protein

(C4BP) and Factor H (FH), facilitated cooperatively by SRS29C and SRS25. SRS57 was also shown to bind FH. Rapid death of Δsrs29CΔsrs25 DKO parasites upon exposure to human serum established that both FH and C4BP are essential for serum resistance. FH conferred a majority of parasite protection by suppressing the alternative pathway amplification loop to abrogate C5b-9 formation and susceptibility to non-immune human serum.

Host sulfated proteoglycans (SPGs) are “self” factors that stabilize FH on host cells to confer protection against complement attack. We hypothesized that an SRS lectin- specific activity for SPGs catalyzes FH recruitment to the parasite surface. The three- dimensional structure of SRS29C alongside biochemical analyses established that SRS29C binds SPGs whereas SRS57 does not, indicating that T. gondii employs both SPG-

iii dependent and -independent FH recruitment mechanisms. We engineered a K16/17S mutation in the basic groove of the SRS29C homodimer that facilitates SPG binding and showed that the mutant failed to bind either SPGs or FH. Transgenic overexpression of

SRS29C in Type I strains enhanced FH recruitment and protected mice from lethal infection by altering inflammatory cytokine production, protecting mice from the cytokine storm induced by Type I strains. Infection with the K16/17S mutant restored virulence, indicating that SPG-dependent FH recruitment appeared necessary to alter induction of the cytokine storm. Type II Dsrs29C parasites were more virulent and surviving mice exhibited greater inflammatory infiltrate in the brain. To establish whether C3 was responsible for these phenotypes, studies using C3-/- mice showed that SRS29C and SRS57 controlled T. gondii virulence in a C3-dependent manner. This work has provided significant mechanistic insight into complement regulation by T. gondii and established the first biological role for an SRS protein expressed on the surface of Toxoplasma.

DEDICATION

This dissertation is dedicated to my husband Juan, to my sisters Marta, Veronika and Agata, to my parents Eugene and Michelle, and to the loving memory of my Babcia, Maria Sienko.

Many thanks, Patricia M. Sikorski

v ACKNOWLEDGEMENTS

I’d like to first and foremost thank my thesis advisor, Dr. Michael Grigg, for his mentorship, support, encouragement and providing me with the opportunity to work on this incredible project.

I would like to thank my colleagues in the Grigg lab, Beth Gregg, Aline Sardinha da Silva, Eliza Carneiro Alves Ferreira, Asis Khan, Viviana Pszenny, Olivia Yanes, Trent Gray, Gloria Adedoyin, Cristina Carvalheiro, Dionne Coker-Robinson, and Andrea Kennard for all of your support, advice, and collaboration. I’d like to especially thank Beth Gregg, for your kindness and positivity, for never giving up on me, and helping me through the ups and the downs. I’m forever grateful for our friendship.

I would like to thank our collaborator Dr. Martin Boulanger at the University of Victoria for the structural work.

To Dr. Chris Hunter, thank you for serving as my external examiner.

I would like to thank the faculty of the Department of Microbiology and Immunology for your continued support for over 10 years, especially Dr. Cole. The department has always felt like home.

I would like to thank my thesis committee, Dr. Calderone, Dr. Fonzi, Dr. Singer and Dr. Sacks for your invaluable insight and guidance throughout this process. I’d like to particularly thank Dr. Calderone, for your continuous support and giving me the opportunity to work in your lab as a master’s student to gain the confidence and skills I needed to begin my career as a researcher.

Many thanks to the Georgetown University-NIH Graduate Partnership Program, especially Allison McBride and Stefan Ambs, for giving me the opportunity, and to my fellow students, especially Thuan, Idalia, and Kinu, for your support and friendship.

A special thank you to Dr. Rembert Pieper, for giving me the opportunity to work on many projects, inspiring me to become a scientist and preparing me for this journey.

To my husband Juan Dominguez, thank you for being exceptionally patient, for being my support system through the long days and weekends in the lab and the long nights writing my thesis. You never complained, always kept me positive at times when I lost hope, and always understood how important this was to me.

I’d like to thank my parents, Eugene and Michelle Sikorski, for their unconditional love and support

To my Sikorski sisters Marta, Veronika and Agata, thank you for being an inspiration to me and never letting me give up.

And lastly, to my late Babcia, Maria Sienko, thank you for teaching me resilience, compassion and the discipline to overcome any obstacle.

Many thanks, Patricia M. Sikorski

vi TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION ...... 1 1.1 The Complement System ...... 1 1.1.1 An ancient immune surveillance system ...... 1 1.1.2 Three major pathways of complement activation ...... 2 1.1.3 Effector functions of the complement system ...... 5 1.1.4 Regulation of the complement system ...... 8 1.1.5 An important bridge to adaptive immune response ...... 12 1.1.6 Intracellular protozoan parasite evasion of the complement system ...... 16 1.2 Toxoplasma gondii ...... 18 1.2.1 The parasite life cycle and transmission ...... 19 1.2.2 Clinical importance ...... 22 1.2.3 Population genetics ...... 24 1.2.4 Acute T. gondii infection in the intermediate host and pathogenesis of infection ...... 27 1.2.5 The immune response to T. gondii and mechanisms of immune evasion ...... 36 1.2.6 Hypothesis, specific aims, and experimental approach ...... 46 1.2.7 Figures ...... 49 CHAPTER 2: SERUM RESISTANCE IN TOXOPLASMA GONDII IS REGULATED BY SURFACE COAT PROTEINS SRS25 AND SRS29C ...... 54 2.1 Introduction ...... 54 2.2 Results ...... 58 2.2.1 Parasite genotype affects C3b deposition but not C3b inactivation or serum resistance ...... 58 2.2.2 Loci associated with regulating C3b deposition are identified by forward genetics ...... 60 2.2.3 Generation of SRSR29C and SRS25 knockout strains ...... 62 2.2.4 SRS29C promotes C3b deposition as a C3 acceptor ...... 63 2.2.5 SRS25 limits C3b deposition ...... 64 2.2.6 Recruitment of soluble lectin and alternative pathway regulators C4BP an FH to the Toxoplasma surface reveals a novel role for tachyzoite surface carbohydrates in lectin pathway activation .... 66 2.2.7 Rapid death of Δsrs25Δsrs29C DKO parasites establishes that FH and C4BP recruitment determine serum resistance ...... 68 2.2.8 Factor H enhances serum resistance by regulating the amplification of the complement response ...... 70 2.2.9 Lack of Δsrs25Δsrs29C DKO parasite killing in mouse serum does not contribute to protection in vivo due to low terminal pathway complement activity ...... 71 2.3 Discussion ...... 72 2.4 Materials and Methods ...... 78 2.4.1 Parasites ...... 78

v 2.4.2 Generation of CZ1Δsrs29C and CZ1 Δsrs25 mutant strains ...... 79 2.4.3 Flow cytometry to determine C3b deposition, Factor H binding and parasite viability ...... 79 2.4.4 Western blotting ...... 80 2.4.5 Factor H blocking and supplementation ...... 81 2.4.6 Mouse serum parasite killing assays ...... 81 2.4.7 Statistical analysis ...... 81 2.5 Figures ...... 82 CHAPTER 3: TOXOPLASMA SURFACE COAT PROTEIN SRS29C RECRUITMENTOF FACTOR H TO EVADE COMPLEMENT AND REGULATE PARASITE VIRULENCE IS SULFATED PROTEOGLYCAN (SPG) DEPENDENT .. 98 3.1 Introduction ...... 98 3.2 Results ...... 101 3.2.1 SRSR29C crystal structure reveals binding groove lined with basic residues at the apical surface ...... 101 3.2.2 An engineered SRS29C dimer specifically binds heparin ...... 103 3.2.3 SRSR29C recruits Factor H through interactions with SPG ...... 105 3.2.4 Trans-over expression of SRSR29C attenuates virulence ...... 107 3.2.5 SRS29C-mediated regulation of acute virulence in the lethal Type I background is SPG-dependent ...... 109 3.2.6 WT SRSR29C expressed in Type I strains alters early cytokine responses ...... 110 3.2.7 SRS29C requires complement protein C3 to regulate virulence in vivo ...... 110 3.2.8 SRS29C alters early cytokine responses in vivo ...... 112 3.2.9 C3 regulates local cytokine responses in vivo and in vitro ...... 113 3.3 Discussion ...... 115 3.4 Materials and Methods ...... 124 3.4.1 Construct engineering, protein production and purification ...... 124 3.4.2 Crystallization and data collection ...... 125 3.4.3 Data processing, structure solution and refinement ...... 125 3.4.4 Sulfated proteoglycan binding studies ...... 126 3.4.5 Parasite culture ...... 126 3.4.6 Plasmid constructs and parasite strains ...... 127 3.4.7 Attachment/invasion assays ...... 127 3.4.8 Flow cytometry to measure C3b deposition and FH recruitment ...... 128 3.4.9 Mouse infections ...... 128 3.4.10 Mouse infection and bioluminescent in vivo imaging ...... 129 3.4.11 Plaque assays to quantify parasite load ...... 129 3.4.12 Ex vivo splenocyte cultures ...... 130 3.4.13 Measuring cytokine production from serum and ex vivo splenocyte cultures by ELISA ...... 130 3.4.14 BMDM culture ...... 130 3.4.15 In vitro infection of BMDMs ...... 130 3.4.16 Luminex assay ...... 131

vi 3.4.17 Measurement of C3a and C5a levels in mouse serum ...... 131 3.4.18 Statistical analysis ...... 131 3.5 Figures ...... 132 CHAPTER 4: TOXOPLASMA SURFACE COAT PROTEIN SRS57 IS COVALENTLY MODIFIED BY COMPLEMENT AND REGULATES PARASITE PATHOGENESIS IN MOUSE MODEL OF ACUTE INFECTION ...... 142 4.1 Introduction ...... 142 4.2 Results ...... 145 4.2.1 Targeted deletion of SRS57 locus in Type I RH and Type II CZ1 strains ...... 145 4.2.2 SRS57 expression impacts C3b and FH binding in vitro ...... 147 4.2.3 SRS57 crystal structure does not support a role for binding SPGs ...... 148 4.2.4 Identification of T. gondii heparin-binding proteins ...... 150 4.2.5 RHΔsrs57 parasites are not deficient in attachment, invasion or extracellular survival kinetics ...... 151 4.2.6 In vivo, RH Δsrs57 infected mice show earlier cachexia and increased parasite burden, but no significant difference in survival ...... 152 4.2.7 Type II Δsrs57 parasites are more virulent and exhibit higher parasite burdens in vivo ...... 153 4.2.8 Protection in vivo is dependent on complement but is not mediated by serum killing of parasites ...... 154 4.2.9 SRS57 regulates systemic and local pro-inflammatory cytokine responses ...... 155 4.3 Discussion ...... 156 4.4 Materials and Methods ...... 165 4.4.1 Parasite strains, host cells and cell culture conditions ...... 165 4.4.2 Targeted disruption and complementation of the SRS57 locus .. 165 4.4.3 Generation of the CZ1Δsrs57 mutant strain using CRISPR/Cas system ...... 167 4.4.4 Flow cytometry ...... 167 4.4.5 Flow cytometry-based complement assays for C3b deposition, FH recruitment, C5b-9 formation and parasite viability ...... 168 4.4.6 Extracellular parasite survival ...... 168 4.4.7 Plating efficiency ...... 169 4.4.8 Rate of replication ...... 169 4.4.9 Mouse infections ...... 170 4.4.10 In vivo imagining ...... 170 4.4.11 Plaque assays to quantify parasite load ...... 170 4.4.12 Ex vivo splenocyte culture ...... 171 4.4.13 Measuring cytokine production from serum and ex vivo splenocytes by ELISA ...... 171 4.4.14 Cloning, expression and purification of SRS57 ...... 171 4.4.15 Crystallization and data collection ...... 172 4.4.16 Data processing, structure solution and refinement ...... 172

vii 4.4.17 Heparin-binding assay ...... 173 4.4.18 SDS-PAGE electrophoresis and Western blot ...... 173 4.5 Figures ...... 175 CHAPER 5: DISCUSSION ...... 190 5.1 Importance of distinct yet cooperative roles for SRS proteins in regulating the human complement system ...... 190 5.2 Implications for the use of murine model for complement studies ...... 199 5.3 Defining novel C3-dependent biological functions of SRS29C and SRS57 in regulating parasite virulence ...... 202 5.4 Implications for complement-mediated lysis independent mechanisms of host protection during acute T. gondii infection and future directions ...... 205 5.5 Figures ...... 217 REFERENCES ...... 221

viii LIST OF FIGURES

Figure 1.1 The complement cascade and effector functions ...... 49 Figure 1.2 T. gondii life cycle ...... 52 Figure 1.3 T. gondii invasion ...... 53 Figure 2.1. Strain-specific difference in C3b deposition does not affect serum resistance in non-immune human serum ...... 82 Figure 2.2. Forward genetics screen and QTL analysis identify several loci regulating strain-specific C3b deposition ...... 84 Figure 2.3. SRS29C promotes C3b deposition and is covalently modified by C3b ...... 85 Figure 2.4. SRS25 limits C3b deposition ...... 86 Figure 2.5. T. gondii recruits alternative pathway regulator Factor H and lectin pathway regulator C4b-binding protein ...... 87 Figure 2.6. Serum resistance requires cooperative recruitment of Factor H and C4b- binding protein by SRS25 and SRS29C ...... 88 Figure 2.7 Factor H recruitment enhances parasite resistance to complement-mediated killing ...... 89 Supplemental Figure 2.1. Titration of NHS determines 10% is optimal concentration for measuring C3b deposition by flow cytometry ...... 90 Supplemental Figure 2.2. C3b deposition on parental strains GT1-SNFR (Type I) and ME49-FUDRR (Type II) ...... 90 Supplemental Figure 2.3. Candidates for reverse genetic studies to validate QTL ...... 91 Supplemental Figure 2.4. Deletion of SRS29C and SRS25 by CRISPR-targeted insertion of HXGPRT gene ...... 92 Supplemental Figure 2.5. SRS25 deletion does not significantly alter expression of SRS29B, SRS29C, SRS34A or SRS57 ...... 93 Supplemental Figure 2.6. T. gondii primarily activates the lectin pathway ...... 94 Supplemental Figure 2.7. Differential surface carbohydrate composition between Type I and Type II strains affects MBL binding and is SRS29C-dependent ...... 95 Supplemental Figure 2.8. SRS25 SRS29C double deletion does not significantly alter SRS29B, SRS34A, or SRS57 expression, but does impact C3b deposition ...... 96 Supplemental Figure 2.9. Low complement activity in mouse serum precludes killing of Dsrs25 Dsrs29C DKO parasites in vivo ...... 97 Figure 3.1. Tandem SRS domains of SRS29C reveal an apical basic surface ...... 132 Figure 3.2. SRS29C dimer dependent binding to heparin localized to basic groove ...... 133 Figure 3.3. SRS29C-mediated FH recruitment is SPG-dependent ...... 134 Figure 3.4. SRS29C trans-overexpression limits wild type parasite proliferation in vivo ...... 135 Figure 3.5. SRS29C alters pathogenesis in lethal Type I infection in SPG-dependent manner ...... 136 Figure 3.6. SRS29C-SPG interaction alters early cytokine responses ...... 137 Figure 3.7. SRS29C-mediated attenuation of virulence in vivo is C3-dependent ...... 137 Figure 3.8. SRS29C expression alters inflammatory cytokine production in vivo ...... 138 Figure 3.9. C3 alters inflammatory cytokine production during acute T. gondii infection ...... 139

ix Supplemental Figure 3.1. SRS29C regulates parasite burden and inflammation in the brains of infected mice ...... 140 Supplemental Figure 3.2. Serum levels of C3a and C5a in C3-/- mice ...... 141 Supplemental Figure 3.3. BMDM infection with serum coated parasites dampens cytokine and chemokine production in bone marrow derived macrophages ...... 141 Figure 4.1. Targeted deletion of the SRS57 gene ...... 175 Figure 4.2. SRS57 expression impacts complement deposition ...... 177 Figure 4.3. Structural divergence of the SRS fold and biological implications of SRS57 crystal structure ...... 179 Figure 4.4. Detection of T. gondii heparin-binding proteins ...... 181 Figure 4.5. In vitro characterization of extracellular survival, attachment/invasion, plating efficiency and intracellular growth of RH Dsrs57 parasites ...... 182 Figure 4.6. In vivo characterization of Type I Dsrs57 parasites ...... 184 Figure 4.7. Mice infected with Type II Dsrs57 parasites exhibit altered pathogenesis .. 185 Figure 4.8. SRS57-mediated protection is dependent on complement but is not mediated by serum killing of parasites ...... 186 Figure 4.9. SRS57 alters systemic and local inflammatory cytokine production ...... 187 Supplemental Figure 4.1. Schematic representation of the strategy for Dsrs57 complementation and expression of GFP-LUC ...... 188 Supplemental Figure 4.2. Extracellular survival of French strains ...... 188 Supplemental Figure 4.3. Paired comparison of extracellular survival between RH WT and French RH WT strains ...... 189 Figure 5.1. Hypothetical model of SRS-mediated regulation of the complement system ...... 217 Figure 5.2. Synopsis of transgenic and deletion mutants and subsequent alterations in interactions with the complement system ...... 219

LIST OF TABLES

Table 1.1 Host regulators of the complement system and their targets ...... 50 Table 1.2 Intracellular protozoan parasite evasion of the complement system ...... 51

xi LIST OF ABBREVIATIONS

AIDS Acquired Immunodeficiency Virus AMA-1 Apical Membrane Antigen 1 AP Alternative Pathway APC Antigen Presenting Cell BCR B cell Receptor C1-INH C1 Esterase Inhibitor C3 Complement Protein C3 C3aR C3a Receptor C4BP C4b-binding Protein C5aR C5a Receptor CCP Complement Control Proteins ConA Concanacalin A CP Classical Pathway CR Complement Receptor CRD Carbohydrate Recognition Domain; Cross Reactive Determinant DAF Decay Accelerating Factor DAMP Damage Associated Molecular Pattern DBA Dolichos biflorus Agglutinin EDTA Ethylenediaminetetraacetic Acid EGTA Ethylene Glycol Tetraacetic Acid ELISA Enzyme Linked Immunosorbent Assay FACS Fluorescent Activated Cell Sorting FDC Follicular Dendritic Cells FH Factor H FI Factor I GalNAc N-acetylgalactosamine GlcNAc N-acetylglucosamine GBP Guanylate Binding Proteins GFP Green Fluorescent Protein GRA Dense Granule Protein HBSS Hanks Buffered Saline Solution HFF Human Foreskin Fibroblasts HIV Human Immunodeficiency Virus IFN Interferon IL Interleukin LP Lectin Pathway LUC Luciferase IDO Indoleamine 2,3-dioxygenase IRG Immunity Related GTPases MAC Membrane Attack Complex MAPK Mitogen Activated Protein Kinase MASP Mannose-Associated Serine Protease MBL Mannose Binding Lectin MIC Microneme

xii MOI Multiplicity of Infection MyD88 Myeloid Differentiation Factor 88 NF-KB Nuclear Factor-kappa B NHS Non-immune Human Serum NK Natural Killer NLR Nod-like Receptor NO Nitric Oxide PAMP Pathogen Associated Molecular Pattern PFA Paraformaldehyde PPR Pattern Recognition Receptor PV Parasitophorous Vacuole PVM Parasitophorous Vacuole Membrane RCA Regulators of Complement Activation RCA I Ricinus communis Agglutinin I ROP Rhoptry Bulb Protein RON Rhoptry Neck Protein SRS SAG1-Related Sequence STAT Signal Transducers and Activators of Transcription Th T helper TLR Toll-like Receptor TCR T cell Receptor TNF Tumor Necrosis Factor WGA Wheat Germ Agglutinin

xiii CHAPTER 1: INTRODUCTION

1.1 The Complement System

1.1.1 An ancient immune surveillance system

At the end of the nineteenth century, a heat labile factor in human blood with bactericidal activity was discovered. It was later termed complement, for its capacity to

“complement” antibody-mediated bacterial lysis (1, 2). To date, this innate serum effector system consists of nearly 50 germline encoded, liver-derived circulating and membrane bound proteins (3, 4). Traditionally, this major arm of the innate immune system functions to detect invading microbes and undergoes a regulated, concerted cascade of events to activate effector proteins that promote inflammation and mediate pathogen destruction.

More recently, broader implications for this system have extended beyond destruction of pathogens and now include removal of debris, stimulation of B and T cell responses, and maintaining homeostasis and intracellular metabolism (5).

A pivotal event in the complement system is the activation of central complement protein C3, which is a member of an evolutionarily old family of secreted thioester containing proteins (TEPs) and plays a key role in innate immunity effector function in both vertebrates and invertebrates (6, 7). Comparative phylogenetic studies have identified complement and complement-like components in most classes of vertebrates (8–10). A C3 homolog in mosquitoes, known as TEP1, has been shown to play an important role in anti- malarial responses (11). Moreover, molecular studies continue to demonstrate the evolutionary origin of the complement system as increasingly ancient; C3 homologues have been identified in sea urchins (12), horseshoe crabs (13), coral (14), and more

1 recently, a gene for a C3-like protein was also identified in the sponge (15). Taken together, evidence for complement-like systems highlight the importance of C3 and C3-like proteins as indispensable, evolutionarily conserved cornerstones of innate immunity (3).

1.1.2 Three major pathways of complement activation

The activation of innate immune responses relies on the detection of highly conserved, invariant microbial structures. The protective function of the complement system in higher vertebrates depends on its capacity to detect invading pathogens, which is achieved through the recognition of pathogen-associated molecule patterns (PAMPS) and damage-associated molecule patterns (DAMPS) via specialized recognition molecules

(16). There are three known pathways for complement activation with distinct mechanisms of recognition. These are known as the classical, lectin, and alternative pathways, which differ in their recognition ligands. Upon binding a particular ligand, conformational changes in the recognition complexes permits autoactivation of pathway-specific proteases and initiates a proteolytic cascade that ultimately leads to the formation of complexes known as C3 convertases (17). All three pathways converge on the activation of central complement protein C3 to generate complement effector proteins C3a and C3b (Figure

1.1)

1.1.2.1 The classical pathway

Initiation of the classical pathway (CP) requires the recognition of antigen-bound

IgG and IgM, or immune complexes via the multimeric C1 complex that consists of the pattern recognition molecule C1q and serine proteases C1s and C1r (18). C1q was first described as a bouquet-like molecule comprised of six N-terminal collagen-like stalks linked to a C-terminal globular head (19). Structural analysis of the globular head indicated

2 that in addition to IgG and IgM, C1q can also recognize non-immune ligands (20–22) and aberrant self-structures to promote clearance of apoptotic cells (23, 24). Recent structural studies determined that the antibody-dependent binding of the C1 recognition complex is dependent on the formation of antigen-bound IgG hexameric structures, which increases the affinity of C1q binding (25, 26). Previous work showed that single IgG binds C1q with lower affinity and this is not sufficient to activate the C1 complex (27). Once C1q-mediated recognition of an antibody-opsonized target occurs, serine proteases C1s and C1r form Ca2+ dependent hetero-tetramers that autoactivate (28). The auto-activation of the C1 complex then allows for a C1s-mediated proteolytic cleavage and activation of C2 and C4, which ultimately form the CP C3 convertase complex, C4bC2a.

1.1.2.2 The lectin pathway

The lectin pathway (LP) is activated in an antibody-independent manner through the recognition of microbial sugars (29, 30). Similar to CP activation, LP multimeric initiation complexes are comprised of recognition molecules and serine proteases. LP recognition of microbial surface carbohydrates relies on several soluble pattern recognition molecules: a superfamily of Ca2+ dependent C-type lectin plasma proteins which include mannose binding lectin (MBL) and collectin-11, and structurally similar ficolins (ficolin-

1, -2, and -3) (31–34). MBL, the best characterized collectin, consists of six trimeric subunits that form a bouquet-like structure, containing a collagenous region as well as carbohydrate recognition domains (CRDs) (35). Ficolins also consist of N-terminal collagen-like domains, however their functional globular domains are fibrinogen-like (31).

Collectins bind a wide range of sugar molecules, such as terminal mannose residues, N- acetyl-D-glucosamine (GlcNAc), whereas ficolins primarily bind acetyl groups such as

3 GlcNAc and N-acetylgalactosamine (GalNAc) (36). The specificity of the ligand binding also depends on the spatial orientation of multivalent, repetitive sugar ligands (37, 38).

MBL and ficolins both associate with MBL-associated serine proteases (MASPs), which autoactivate upon recognition of microbial oligosaccharides, and mediate the cleavage of

C2 and C4 to form a C4bC2a C3 convertase complex (31, 32).

1.1.2.3 The alternative pathway

The alternative pathway (AP) is evolutionarily the oldest and most conserved pathway of the complement system. The AP, also once known as the properdin pathway, is an antibody-independent pathway that does not rely on recognition for activation (2, 39).

Rather, its initiation relies on the failure to regulate low-level spontaneous hydrolysis of

C3 into C3(H2O), an event also known as “tick-over”. This hydrolysis results in a conformational rearrangement to expose the C3 internal thioester, which permits the formation of a soluble AP C3 convertase (C3b(H2O)Bb) after interacting with Factor B and

Factor D (40, 41). This complex cleaves C3 and generates a nascent C3b activated protein that can covalently bind to pathogen surfaces (42–45). These interactions are stabilized by positive regulator properdin and require Mg2+ (46). Properdin has also been identified as a recognition molecule for AP activation (47, 48), however this remains controversial and has recently been challenged due to its dependence on initial C3b deposition (49). Rather,

AP is indiscriminate in that C3b arbitrarily probes foreign and self surfaces alike, which must be stringently regulated by pre-formed regulators. Thus, the ability of the AP to discriminate between self and non-self relies on both membrane-bound and soluble regulatory molecules that associate with host cell surfaces by recognizing “self “molecules such as sialic acid and glycosaminoglycans (50, 51). Foreign cells, which lack these

4 surface-associated regulators, are not able to control AP activation. Arguably, the most important function of the AP is its capacity to use C3b generated by both classical and lectin pathways as a platform for forming new AP C3 convertases, establishing a positive feedback loop that rapidly amplifies the complement response (52, 53). Several studies have shown AP-mediated amplification can account for up to 80-90% of complement activation in IgG induced CP and mannan induced LP activation models (54–57).

1.1.2.4 All pathways converge at C3

All three pathways converge on producing the same effector molecules through cleavage of C3 by pathway-specific C3 convertases. C3, the most abundant complement protein in plasma (~1.0ug/ml), is a 187 kDa protein consisting of a stable 75 kDa beta chain and a 115 kDa thioester containing alpha chain, held together by a disulfide bond (58, 59).

During activation of C3, C3a, a soluble inflammatory peptide, is released whereas the larger fragment, C3b, covalently couples to host and pathogen surfaces unless an AP regulator is present to inactivate C3b. A conformational change upon C3 cleavage exposes a reactive thioester that allows C3b to react with nearby hydroxyls and amines (42, 44, 58,

60–62). Both C3a and C3b have critical roles in mediating the antimicrobial effector functions of the complement system, these are described more fully in sections 1.1.3.2 and

1.1.3.3.

1.1.3 Effector functions of the complement system

Complement activation contributes to immune defense by generating numerous biologically active molecules, which include anaphylatoxins (C3a, C5a), opsonins (C3b, iC3b, C3dg, C3d), and terminal pore formation (C5b-9 or membrane attack complex) (63), that have several important protective functions.

5 1.1.3.1 Membrane attack complex formation and direct lysis

The most direct effector function of the complement system is mediated by the terminal pathway, which concludes in the assembly of the membrane attack complex

(MAC) that disrupts cell membranes and leads to cell lysis (64). MAC, also known as

C5b-9, is composed of several complement proteins: C5, C6, C7, C8 and C9 (65, 66). C3b is a fundamental component in the formation of a C5 convertase to mediate the last enzymatic step of the complement cascade and first step in the terminal pathway of the complement cascade. The first step of the terminal pathway requires C5 cleavage by C5 convertase, which is formed when additional C3b molecules associate with existing C3 convertases (67). High densities of C3b are required to drive the enzymatic activity of this complex towards preferentially cleaving C5 (68, 69) into C5a, a soluble anaphylatoxin, and

C5b. C5b initiates pore assembly, which occurs in distinct stages of intermediate formations that progressively insert into the cell membrane. C5 cleavages exposes a C6 binding site on C5b (70). Subsequent association of C7 with C5bC6 increases affinity for lipids, allowing for association of the C5b-7 complex with the phospholipid membrane bilayer (71–73). C8 binding to this complex induces membrane insertion (74, 75), and allows for C9 to bind, polymerize and form a stable fully-formed MAC (64, 76, 77). Lytic function is attributed to alterations in membrane stability (64, 78, 79) and Ca2+ influx (80).

Notably, many pathogens are resistant to MAC formation, thus complement activation overcomes this limitation by multiple effector molecules to ensure additional layers of immune defense (81).

6 1.1.3.2 Opsonin-mediated phagocytosis

C3b and its catabolites (iC3b, C3dg) are versatile effector molecules that function as the major opsonins in the absence of antigen-specific antibodies to help mediate phagocytosis and pathogen clearance. Several families of complement receptors (CRs) on myeloid cells recognize these opsonins, resulting in complement-mediated phagocytosis

(82, 83). CR1, a short consensus repeat module containing protein, is expressed on monocytes, neutrophils, and erythrocytes, and binds with high affinity to C3b but can also bind iC3b, C3dg and C1q to facilitate phagocytosis, removal of erythrocytes, and immune complexes (84–86). CR3 (CD11b/CD18) and CR4 (CD11c/CD18) are phagocytic receptors belonging to a versatile family of β2 intergins that consist of transmembrane heterodimers composed of an α chain (CD11b, CD11c) and common β chain (CD18) (87), both of which share iC3b as a ligand. Lastly, an immunoglobulin-type receptor CRIg is expressed on liver macrophages, known as Kupffer cells, and has recently been shown to mediate clearance of circulating pathogens through interactions with C3b and iC3b (88).

CR2 is another complement receptor expressed on B cells and follicular dendritic cells that interacts with C3b catabolite C3dg and plays a role in stimulating B cell responses. This complement receptor will be discussed more fully in section 1.5.1.

1.1.3.3 Anaphylatoxins C3a and C5a are pro-inflammatory mediators

Additional effector functions of the complement system rely on the interaction of complement effector proteins with cognate receptors that are expressed on both immune and non-immune cells. Anaphylatoxins C3a and C5a are small 10 kDa, soluble, pro- inflammatory polypeptides released upon C3 and C5 activation. These molecules are important danger signals that exert their biological function upon binding their cognate

7 receptors, C3aR and C5aR, that are expressed on a variety of cells. Signaling through these receptors induces a wide range of activities, which include chemotaxis, oxidative burst, release of histamine from basophils and mast cells, immune cell activation, vasodilation, and induction of cytokine responses (89–94).

1.1.4 Regulation of the complement system

The activation of the complement system is unfortunately not limited exclusively to pathogens. Due to the non-discriminatory nature of, in particular, the AP and its capacity to generate a rapidly amplifying positive feedback loop, complement can activate on bystander cells. Thus, the host must stringently regulate the complement system to prevent harm to the local environment and prevent excessive inflammation. Expression of several membrane bound and soluble complement regulatory proteins achieve protection from inadvertent complement activation on host cells. These regulators target all major points within the cascade, including pathway-specific initiation events, activation events mediated by C3 and C5 convertases, and effector steps such as MAC formation (Table

1.1). The importance of complement regulators is highlighted by several diseases, such as hemolytic uremic syndrome (aHUS) and age-related macular degeneration, which are associated with mutations and polymorphisms in soluble regulator Factor H that diminish its regulatory function, leading to uncontrolled complement activation on host tissues and organ specific damage (95, 96).

1.1.4.1 Initiation regulators

Classical and lectin pathways rely on distinct pattern recognition complexes, thus there are several soluble regulators that specifically target these initiation events. These types of soluble inhibitors specifically target the activation of serine proteases that are

8 complexed with the classical and lectin pathway recognition molecules. The only known regulator of the CP is C1 esterase inhibitor (C1-IHN), a soluble serpin (serine protease inhibitor) that inactivates the C1 complex by binding the catalytic sites of proteases C1r and C1s (97). C1-INH further dissociates the inhibited protease complexes from both the

C1q molecule and IgG activating surfaces (97, 98) and binds irreversibly (99). C1-INH has also been shown to inhibit the LP by inhibiting serine proteases MASP-1 and MASP-2

(100). Additional LP MASP inhibitors include MAp44 and MAp19 (101–103). By regulating the initiation step, C2 and C4 activation is limited and thus precludes downstream C3 convertase (C4b2a) assembly.

1.1.4.2 Convertase regulators

C3 and C5 convertases are common targets for regulators because of their gatekeeper role in driving complement activation, amplification and effector generation.

Convertase directed regulators consist of both membrane-bound and soluble regulators.

The strategies to control convertase formation and its activity are multifaceted and include destabilization of convertase complexes, decay accelerating activity of the catalytic domain, and co-factor activity for serine protease Factor I to degrade C4b and C3b and eliminate the possibility of these effectors to build fresh convertases (104). C4b-binding protein (C4BP) is a CP and LP regulator that targets the C4b component of the C4bC2a convertase. C4BP is part of a family of complement control proteins (CCPs) known as regulators of complement activation (RCA), which are defined by short consensus repeats

(SCRs), ability to bind complement proteins C4b or C3b, and their clustering on

Chromosome 1 at the q3.2 locus (105, 106). C4BP protein has an unusual spider-like structure (107), consisting of 7 identical a chains, each containing 8 SCRs and a single b

9 chain of 3 SCRs. C4BP targets the C3 convertase by accelerating its decay (108) and preventing the formation of new convertases by acting as a co-factor for the proteolysis of

C4b (109). Other RCA proteins that regulate C3 convertases are membrane bound regulators decay accelerating factor (DAF; CD55), CD46 (also known as MCP, membrane cofactor protein), and complement receptor 1 (CR1) that similarly accelerate the decay of

C3 convertases and act as cofactors for Factor I (Table 1.1) (106).

In contrast to CP and LP activation, which occurs in response to pathogens or danger signals, the low-level constitutive activation, or “tick-over”, of the AP has the potential to indiscriminately bind both foreign and self surfaces alike, and subsequently rapidly amplify if left uncontrolled. Thus, the AP is stringently and predominately regulated by soluble regulator Factor H (FH), formerly known as b1H. FH is also part of the RCA family, and consists of 20 homologous complement control proteins (CCPs), or

SCRs, that give it the appearance of a flexible “beads on a string” that can fold back on itself (110). Like the previously described RCA proteins, FH has decay accelerating activity of the AP convertase by dissociating Bb (111, 112) and co-factor activity for inactivating C3b, which plays a critical role in regulating the amplification loop.

1.1.4.3 Terminal pathway regulation

Regulators of the terminal pathway, which include vitronectin (also known as S protein) and clusterin, which primarily bind or scavenge components or intermediary pore complexes (C5b-7, C5b-8, C5b-9) to block their incorporation into host cell membranes

(113) or prevent the polymerization of C9 (114), a key step in MAC assembly. CD59, or protectin, is a membrane bound regulator that binds C8 and C9 within C5b-8 and C5b-9, respectively (115–118).

10 1.1.4.4 Discrimination of self-versus non-self

The AP, unlike the CP and LP, does not recognize any particular pattern in order to activate. Rather, low level spontaneous hydrolysis of C3 leads to production of C3b that is constantly probing cells and randomly depositing on any surface (50). Thus, AP activation occurs by default and relies on the presence of host regulators to distinguish between activating versus non-activating surfaces. The discrimination between self and non-self surfaces is actually performed by soluble regulator FH. Early on, evidence suggested that

Factor H was equipped with self-homing mechanisms that bias complement regulation on self-surfaces over foreign ones (104).

The first evidence for a mechanism of host cell protection from AP originated from a study that showed protection from AP activation on erythrocytes was lost after the removal of surface sialic acid from its surface (112, 119), indicating that polyanions were used by FH to identify self surfaces. Later, another host polyanion heparan sulfate, a type of glycosaminoglycan chain found on host sulfated proteoglycans (SPGs), was later shown to inhibit AP activation and interact specifically with FH (120, 121). Notably, the interactions between host polyanions and FH was shown to increase the affinity of FH to

C3b. This affinity was found to be largely dependent on the context in which C3b was engaged with its target (121, 122). High affinity of FH for C3b was favored on C3b coated surfaces with sialic acid or SPGs (112), resulting in the formation of a surface bound tripartite complex of SPG, C3b, and FH . On the contrary, non-activating surfaces lacking

“self” molecules have low affinity for FH and can bind Factor B to generate AP convertases. Furthermore, in the absence of HSPGs or sialic acid, such surfaces targeted

11 by the CP or LP, C3b complexed with C4b or IgG is protected from FH binding and inactivation (123).

Structural and functional studies identified several binding sites for both SPGs and

C3b that allows it to bind both molecules. Of the 20 complement control protein domains

(CCPs) that make up FH, domains 1-4 have been attributed to C3b binding, cofactor activity and decay accelerating activity (124, 125), whereas domains 7-8 and 19-20 mediate host specific interactions (126–128). However, several studies have also shown that domains 19-20 bind C3b, iC3b and C3d, thus overlapping with polyanion binding sites

(129). Recently, structural analysis has resolved this conundrum at the molecular level, demonstrating that the CCP domains 19 and 20 simultaneously bind C3d and surface glycosaminoglycans, respectively (130). Thus, polyanions and C3d are two sole distinguishing features of non-activator surfaces that are recognized by FH, leading to the breakdown of C3b. One study has also demonstrated that C4BP (131) also requires recognition of heparin to discriminate between self and non-self surfaces. However, the molecular and structural basis for the homing of C4BP to self surfaces is not nearly as well characterized as FH.

1.1.5 An important bridge to adaptive immune response

Although the complement system has traditionally been defined as an innate serum effector system, recent paradigm-shifting observations have emerged, revealing some unexpected novel functions for the complement system. Most notably, complement has been implicated in shaping the adaptive immune response. Complement activation products have recently been added to the list of factors critical for instructing both B cells and T cell responses. Anaphylatoxin receptors and complement receptors on lymphocytes

12 interacting with complement activation products have been shown to play a role in activating, differentiating, and enhancing B and T cell responses (132–134). Importantly, these studies have also challenged the paradigm that complement solely impacts extracellular spaces, as cellular sources and intracellularly activated complement have recently been described.

In contrast to innate immunity, the adaptive immune response is highly antigen- specific and requires time to develop a defined response. The development of these responses relies heavily on signals and information acquired from the innate immune system to drive a particular response against an invading pathogen (135). Priming B and T cell responses requires antigen presentation by antigen presenting cells that engage B cell receptors (BCRs) and T cell receptors (TCRs), interaction with costimulatory molecules, and production of soluble factors that stimulate differentiation of specialized lymphocyte subsets. Additionally, the specialized environments of lymphoid organs facilitate these intimate interactions. Complement activation products and their receptors have recently been shown to play important roles in these events.

1.1.5.1 Regulation of B cell responses

B cells mediate their protective function by generating antigen-specific antibodies.

Antibody production by B cells require the right stimuli and signals to allow for B cell maturation, which includes presentation of antigen and receptors, as well as a specialized environment of the peripheral lymphoid organ. Optimal activation of B cells requires both antigen recognition by the B cell receptor (BCR), as well as colligation with co-receptors to amplify signaling events. A role for complement in the adaptive humoral response was first described when complement receptors were identified on B cells (136). Later,

13 observations of decreased antibody titers and smaller germinal centers in mice lacking C3 or CR1/2 indicated that complement played a specific role in humoral immunity (137–

140). The role of CR2 was elucidated when it was discovered to be a co-receptor on B cells that cooperates with the BCR and CD19 to efficiently recognize C3d-covalently bound antigen (141, 142). Furthermore, C3d coupled antigens have been shown to enhance immunogenicity of antigen by lower the threshold for B cell activation to generate an antibody response (142, 143). Taken together, these studies demonstrate that antigen covalently linked to C3d promotes the cross linking of the BCR with CD19/CR2 co- receptors to enhance B cell immunity (144).

These critical interactions between complement, antigen, and B cells takes place in the germinal centers (GCs) of peripheral lymphoid organs (145, 146) and are dependent on follicular dendritic cells (FDCs), a specialized type of dendritic cell that resides in the B cell follicle of GCs. CR2 is also expressed on follicular dendritic cells (FDCs) (84, 147).

FDCs trap C3d-tagged immune complexes transferred from follicular B cells in a CR2- dependent manner (148, 149) and retain native antigen for long periods of time. Without

CR2, GCs and B cells fail to mature and undergo death (146), thus demonstrating CR2 also plays a role in GC development.

1.1.5.2 Regulation of T cell responses

The discovery of extra-hepatic synthesis of complement proteins suggested that local cellular sources of complement production may have special functions outside of its traditional roles. Studies in both murine and human cells demonstrated that many cell types, including epithelial cells, myeloid cells, and lymphocytes can produce and activate their

14 own complement (150–154). Recently, cell-derived complement components have been shown to play a critical role in promoting T cell activation and differentiation (155).

Upon antigen stimulation, T cells require costimulatory signals to activate and proliferate. During cognate interactions, both APCs and CD4+ T cells surprisingly upregulate alternative pathway components C3, C5, Factor B and Factor D (152).

Furthermore, these cells also upregulate expression of anaphylatoxin receptors C3aR and

C5aR (156). The local activation and secretion of these anaphylatoxins were shown to act in an autocrine and paracrine fashion as co-stimulatory signals to promote cytokine production, T cell immunity, and T cell expansion. (152, 157). Blocking anaphylatoxin engagement with cognate receptors using C3aR and C5aR antagonists prevented upregulation of IL-12 mRNA in dendritic cells and IFN-g production in responding CD4+

T cells (152). Importantly, these studies determined that C3aR and C5aR had overlapping and partially redundant functions in the murine model and required deficiency or blocking of both receptors to completely abrogate T cell responses.

Similarly, upregulation of locally produced complement proteins was also observed in activated human CD4+ T cells. Recently, it has been determined that CD4+ T cells have intracellular stores of C3 and C5 that are cleaved by cathepsin-L to produce C3a and C5a, rather than by traditional convertases, upon TCR activation (150). Upon T cell activation, this intracellular activation system is shuttled to the cell surface, where C3aR and CD46 engage C3a and C3b, respectively. The C3 activation products C3a and C3b generated by human T cells drive Th1 differentiation and IFN-g production (158). At steady state, C3a can also bind intracellular C3aR and maintain T cell viability by sustaining basal mTOR activation (150). Importantly, these T cell-modulating functions are independent of serum

15 circulating complement and occur in serum free conditions. Uncontrolled intracellular C3 activation in CD4+ T cells have been shown to contribute to increased Th1 responses associated with several autoimmune diseases (150). Taken together, these studies are groundbreaking in that they identified sources of complement that are not liver-derived or extracellular and function endogenously in regulating T cell responses.

1.1.6 Intracellular protozoan parasite evasion of the complement system

The complement system is the first line of defense against many pathogens, including intracellular protozoan pathogens. However, this group of pathogens has evolved sophisticated strategies to evade complement attack to establish infection and cause disease. These evasion mechanisms fall into two general categories: parasite encoded regulators that often mimic the function of host regulators and acquisition of host regulators

(Table 1.2). Often, these parasites employ multiple surface factors to target several points within the cascade to confer protection. Several parasite factors involved in complement regulation for intracellular protozoan pathogens Trypanosoma cruzi, Leishmania, and

Plasmodium have been identified, however these factors are completely unknown for other protozoan parasites, including Toxoplasma gondii.

1.1.6.1 Parasite-specific complement regulators

The infective stage trypomastigotes of Trypanosoma cruzi, the etiological agent of

Chagas disease, is resistant to complement mediated lysis by expressing several surface proteins that regulate C3 convertase formation. Complement regulatory protein (CRP, gp160) binds C3b and C4b to dissociate the classical and alternative C3 convertases (159).

Complement C3 receptor inhibitor transpanning (CRIT) also regulates both classical and lectin pathways by binding both C1q and MBL (160, 161). Trypomastigote decay

16 accelerating factor (T-DAF) shares 40% sequence homology with human DAF and is functionally analogous to its human counterpart by interfering with C3 convertase formation (162).

Leishmania also evades and exploits the complement system by encoding its own set of parasite specific factors. GP63 is a parasite glycoprotein that is highly conserved among Leishmania species and functions as a surface protease the binds C3b and inactivates it (163, 164). LPG is also another major surface component that is involved in the resistance of terminal pathway activation by releasing C5b-9 complexes (165–167).

1.1.6.2 Acquisition of host factors

Plasmodium, the causative agent of malaria, is resistant to both human and mosquito complement and complement-like systems. Recent studies have shown two soluble host regulators Factor H and C1-INH are acquired by blood stage merozoite surface proteins Pf92 and PfMSP3.1, respectively, however the precise mechanisms remain unknown (168, 169). In the mosquito stages, Plasmodium protein GAP50 has also been implicated in binding FH (170). Surface protein Pfs47 evades the mosquito complement like system protein TEP1 by a distinct mechanism that does not rely on host regulators, but rather suppresses midgut nitration responses that are critical for activating the complement- like system (171, 172).

Toxoplasma gondii is arguably the world’s most successful parasite. It can infect all mammals and up to a third of the world’s human population are infected. An early study established that T. gondii is resistant to complement mediated lysis in non-immune serum by inactivating C3b (173). Surprisingly, since this original discovery in 1989, no further work has been done to identify the parasite factors that interact with complement or the

17 molecular basis for complement resistance. Considering the elaborate strategies utilized by several intracellular protozoan pathogens described above to inactivate the complement pathway, it seems likely that T. gondii also employs several surface proteins to overcome this important first line of defense. Toxoplasma gondii is a complex organism whose success depends on its lifecycle, virulence factors, and immune evasion strategies. To gain a better understanding of this organism and its interactions with the immune system, an overview of Toxoplasma biology and pathogenesis is presented in the next section.

1.2 Toxoplasma gondii

Toxoplasma gondii is a ubiquitous, obligate intracellular eukaryotic parasite within the phylum Apicomplexa. Apicomplexan parasites are amongst the most diverse and prevalent group of single-celled pathogens in the world, consisting of more than 6,000 species (174) that infect almost all mammal species. These parasites are unified by a distinct, highly polarized morphological feature at their apical end that consists of a sophisticated network of secretory organelles. Many of the parasites in this phylum are of clinical and veterinary importance, which include Plasmodium, Cryptosporidium,

Neospora, Babesia, and Sarcocystis. These parasites have complex life cycles consisting of stage- and often host- specific forms, which provides a challenge for studying their biology. However, of all the species, T. gondii is arguably the most experimentally tractable. Although all stages of T. gondii cannot be grown in vitro, the pathogenic asexual tachyzoite stage is easily grown, maintained, and manipulated in the laboratory and the chronic stage can be maintained in animal models. Well established methodologies for forward (175, 176) and reverse genetics (177–181) and animal models have provided

18 significant advantages for studying the biology of this parasite (182). Therefore, it is an ideal model organism to study shared aspects of Apicomplexan biology, which include invasion, host cell modifications, and host-pathogen interactions.

1.2.1 The parasite life cycle and transmission

Apicomplexan parasites have complex life cycles with vast host specificities, which can range from a single host to most mammals. T. gondii is a successful parasite that is capable of infecting essentially any nucleated cell in any mammal (183). It belongs to a group of enteric, tissue encysting parasites, known as the coccidia. All coccidian parasites are intestinal parasites that have an asexual and sexual cycle (184). T. gondii is an exceptional coccidian parasite in that its transmission is not restricted to either a single host species or is exclusively fecal-oral. Unlike Cryptosporidium, Eimeria, or Sarcocystis which cannot propagate between intermediate hosts (185), Toxoplasma can be propagated both sexually in its definitive host or asexually by carnivory, because tissue cysts in all mammals are orally infectious, so it does not need to undergo its sexual cycle to be transmitted in nature (Figure 1.2). It is this highly flexible lifecycle that has significant implications for its transmission, genotypic diversity, expansion into new hosts, and virulence across its large host range.

The T. gondii sexual cycle occurs in the gut of its definitive host, which includes all felids, and results in the shedding of a large number (often in excess of 1x10e9) of highly infectious oocysts that are highly stable and can persist in the environment for 6 months to 1 year and remain infectious (186, 187). T. gondii can be acquired by intermediate hosts through the fecal-oral route, carnivorism, and vertically from mother to fetus. In humans, infection is acquired by the ingestion of either infected meat or oocysts

19 contaminating water and food. Both of these life cycle forms are orally infectious.

Sporozoites are the infections forms present in oocysts whereas bradyzoites are the infectious form present in tissue cysts (188). The intake of oocysts contaminating water or vegetation or tissue cysts by eating an infected animal ensures transmission to both herbivores and carnivores. Inside the intermediate host, the parasite undergoes two phases of asexual development.

1.2.1.1 Asexual stage

Once oocysts or tissue cysts are ingested, parasites excyst in the small intestines, invade the intestinal epithelium and differentiate into the rapidly replicating asexual form, known as the tachyzoite (189). The tachyzoite mediates acute infection in the intermediate host by rapidly replicating through endodyogeny and disseminates systemically via the blood and lymph to distal organs (189). This parasite stage is also capable of crossing biological barriers, including the blood-brain, the blood-eye barriers as well as the placenta, the latter resulting in vertical transmission of infection from mother to fetus. During acute infection, tachyzoites induce a strong inflammatory response, where most of the parasites are effectively cleared by the immune system. Ultimately, in vitro studies suggest that immune pressure and other unknown factors drive the parasites to differentiate into the second transmissible asexual form, the semi-dormant bradyzoites that are contained within a tissue cyst (190–192). Bradyzoites are known to molt and express a new suite of surface antigens, which are thought to prevent parasite clearance by escaping anti-tachyzoite immunity and promoting cyst persistence for the life-time of the host (193–195). Under several immunologically driven circumstances, bradyzoites can transition back to tachyzoites and thus reactivate acute infection.

20 Both asexual forms are critical to the parasite’s success. Tachyzoites contribute to the rapid replication and dissemination of the parasite across several biological barriers, which are important prerequisites for chronic infection. The bradyzoite stage is a critical factor in Toxoplasma pathogenesis, persistence and transmission (196). However, it’s major contribution to T. gondii’s success extends beyond its capacity to be orally infective, as it is capable of persisting in the presence of a functional immune system for the life of the host. Notably, current treatments for toxoplasmosis are only effective against the tachyzoite stage (197). Bradyzoites appear to be impervious to both the immune system and drugs, which facilitates the parasite’s lifelong persistence and ability to propagate in nature.

1.2.1.2 Sexual stage

The definitive host is also infected by ingesting either oocysts from the environment or by eating infected prey. Once ingested tissue cysts or oocyst excyst in the gut, bradyzoites and sporozoites invade feline enterocytes and undergo a developmental transition to merozoites, rather than tachyzoites. Merozoites then divide to amplify the infection before a developmental cue initiates their terminal differentiation into either micro- or macro-gametocytes (male and female gametes, respectively) that fuse to produce a diploid zygote (198–200). The fertilized zygote then undergoes several rounds of nuclear division to produce an 8n oocyst. Oocyst production peaks 5-8 days post infection and are then shed in the feces in numbers as high as 100 million (198). When shed into the environment, oocysts are unsporulated and do not become infectious until they sporulate, a process which requires oxygen and moist, slightly acid conditions. Meiosis occurs during sporulation and each oocyst produces two sporocysts, with each sporocyst containing four

21 infectious haploid sporozoites (201–203). Oocysts are extremely durable and resistant to environmental and chemical destruction, however, they are sensitive to desiccation and high temperatures (204–207).

The biological significance of the sexual cycle is two-fold: widespread environmental transmission and genetic diversity. The generation of a large number of highly infectious oocysts in just one shedding exponentially potentiates environmental transmission. Furthermore, environmental transmission due to feline shedding in close proximity to water supplies or periods of high runoff has caused several waterborne outbreaks (207–210). The sexual cycle is incredibly fecund, with one cat generating as many as 100 million recombinant progeny (211), which can reshuffle parental alleles into

100 million new combinations and such “mixis” of genetic material produces new strains that can be released into the environment. Together, these factors shape the population structure and have been shown to significantly impact virulence across various hosts (212,

213).

1.2.2 Clinical importance

T. gondii infection is a life-long, chronic infection due to the persistence of tissue cysts. It is estimated that one-third of the world’s population is chronically infected with

T. gondii, however, prevalence is highly variable depending on geographical location. For instance, in the United States, seroprevalence ranges from 10-20%, whereas in endemic areas like France and Brazil, prevalence can be as high as 50% and 80%, respectively (214).

In healthy individuals, acute infection is typically asymptomatic, although some individuals present with flu-like symptoms, including fever, headache, muscle pain and enlarged lymph nodes (215). However, there are groups of individuals that are at high risk

22 for developing a severe infection, which include congenitally infected fetuses, newborns, and immunosuppressed individuals. In rare cases, healthy individuals may present with acute acquired disease. Toxoplasma infection can affect multiple tissues, but the most common clinical manifestations of toxoplasmosis involve the brain and eye. Thus, a critical factor underlaying the pathogenesis of toxoplasmosis is the parasite’s ability to disseminate and cross various biological barriers during acute infection.

1.2.2.1 Congenital toxoplasmosis

Congenital toxoplasmosis is the result of primary infection during pregnancy.

Transplacental passage of tachyzoites can occur during acute infection and result in infection of the fetus (216). The annual global incidence of congenital toxoplasmosis is estimated to be 190,00 cases (217). The severity of disease depends on which trimester in the pregnancy infection occurs, with a large range of symptomatic to asymptomatic manifestations of the disease that can affect the fetus (218). Infection during the first trimester often results in more severe outcomes, which include spontaneous abortion, hydrocephaly, and severe neurological effects, whereas transmission during the end of the pregnancy results in typically asymptomatic infection (215, 219). Although the incidence of symptomatic infections at birth are low, the majority of asymptomatic infants infected congenitally will present with sequalae later in life. For example, greater than 75% of those born with asymptomatic disease at birth present with recurrent chorioretinitis later in life

(220).

1.2.2.2 Immunodeficiency and toxoplasmosis

Toxoplasmosis in immunodeficient individuals is the result of reactivation of a chronic infection. Chronically infected individuals who become immunosuppressed due

23 HIV/AIDs, chemotherapy, or organ transplant recipients are at risk for developing reactivated systemic toxoplasmosis. The importance of cell mediated immunity in controlling T. gondii infection was underscored by the 1980s AIDS epidemic, as T. gondii infection was the most common opportunistic infection of the central nervous system that lead to lethal encephalitis when left untreated (221). Toxoplasmic encephalitis in these patients occurred in about 10-50% of seropositive patients, depending on geographical location (222).

1.2.2.3 Acquired ocular toxoplasmosis

T. gondii infection is the most common cause of infectious uveitis in healthy adults worldwide. In the U.S., ocular toxoplasmosis accounts for 1-2% of eye disease (223).

Ocular toxoplasmosis often presents as a focal lesion in the anterior of the eye and is typically a recurrent disease with associated inflammation, necrosis, and some degree of visual impairment (215, 224). Ocular toxoplasmosis in South America accounts for up to

20% of eye disease in regions of Brazil and often is more severe due to the presence of more virulent genotypes circulating in those regions (225–227).

1.2.3 Population genetics

Although considered a single species, T. gondii population genetics is both genetically diverse and highly clonal depending on the geography and host species sampled. Early molecular typing studies of strains from symptomatic humans and livestock from Europe and North America identified a surprisingly limited, clonal T. gondii population structure (228, 229). For a parasite with a fecund sexual cycle, the population genetic structure was unexpectedly limited to essentially three predominant genotypes referred to as Type I, II and III in these sampled niches (230). Molecular analyses

24 determined that these strains were highly similar and shared a common ancestor (231). In striking contrast, the majority of strains isolated from South America were non-clonal, atypical strains indicating the population structure was quite diverse in this geographic region (232, 233). More in depth analyses determined that South American strains are comprised of distinct lineages that are not found in North America (231). With the advent of whole genome sequencing, greater resolution of the differences found among 62 circulating strains, thought to encompass total genetic diversity within the species, revealed that the population genetic structure was comprised of 6 major clades (or races) and that the majority of strains resolved into 16 haplogroups, with each haplogroup having a distinct geographical distribution(234).

This discrepancy in geographical diversity can be explained by the parasite life cycle. Clonal expansion of strains in North America and Europe suggests propagation of asexual tissue cysts predominantly through carnivory and successive transmission through intermediate hosts (188, 230). Alternatively, since the rate of co-infection of more than one strain in cats may be a rare occurrence, clonal expansion by sexual inbreeding can likewise result in progeny that appear to be identical to the parental strain (211, 235). In contrast, the highly diverse population structure in South America has been attributed to a robust sexual cycle with a high degree of outcrossing (231, 232, 235). Taken together, the population structure across several geographical regions has been shaped by both sexual and asexual propagation of this parasite, however it is unclear what factors have contributed to these regional differences in genetic diversity.

25 1.2.3.1 Implications of parasite genotype on virulence and disease

The three main clonotypes only differ by 1-2% at the nucleotide level (212).

However, despite this seemingly little variation, parasite genotypes can have vastly different phenotypes. Strain-specific differences in parasite growth, dissemination, virulence and induction in innate immunity have been observed (236, 237). In fact, the best example of these differences is virulence in laboratory mice. The archetypal Type I, II and

III strains were initially designated as such based on their virulence in mice, where Type I

3 strains are uniformly lethal at low doses (LD100=1), and Type II (LD50=10 ) and Type III

5 (LD50=10 ) strains are avirulent at low dose inoculums and establish chronic, non-sterile persistent infections (228, 238). Experimental crosses have demonstrated that a single cross between two avirulent strains in this haploid parasite (e.g., Type II x Type III) can yield progeny that are highly virulent in mice (212). Hence, sexual recombination, by its ability to reshuffle genetic diversity, can bring mouse virulent alleles that are epistatic in both parental genetic backgrounds into new combinations that alter the biological potential of the progeny. Thus, the molecular basis for Toxoplasma virulence in mice can be attributed to specific combinations of parasite polymorphic effector proteins that function to inactivate host immune signaling networks to alter the parasite’s pathogenesis (239, 240).

The contribution of genotype to human disease is less defined. There are some reported associations of disease manifestations with strain type. While all three clonal lineages are found in human infection, the majority of human toxoplasmosis and congenital infection in North America and Europe is associated with Type II strains (229). Infections with Type I strains are less common, however these strains have been associated with groups of immunocompromised patients and Type III strains are primarily found in

26 livestock and are not commonly found infecting humans (228, 241). Human disease is also highly dependent on geographical location, where atypical strains have been associated with more severe congenital infections in Brazil (229, 242, 243). Though rare in the U.S, the severity of eye disease in patients with ocular toxoplasmosis has been attributed to more diverse, atypical strains found in South America (226, 244, 245).

1.2.4 Acute T. gondii infection in the intermediate host and pathogenesis of infection

Acute disease in the intermediate host is mediated by the tachyzoite stage, which is able to actively invade, rapidly replicate, and disseminate through the host. The ability of tachyzoites to cross biological barriers and infect many different tissues significantly contributes to T. gondii pathogenesis and persistence. To achieve this, the parasite employs several strategies that rely on various parasite structures, organelles, and proteins that work together to coordinate motility, invasion, and the establishment of an intracellular replicative niche. Although the immune response is critical in controlling parasite replication, it is unable to clear the infection. It has become increasingly clear over the past two decades of research that Toxoplasma encodes large suites of highly polymorphic effector proteins that it uses to evade and/or modulate host immune responses to facilitate infection. Thus, understanding the coordinated efforts of these parasite-specific factors and strategies to influence disease outcome and parasite persistence is of critical importance.

Contributing to the complexity of this parasite, several observed differences between human and murine immune responses to T. gondii indicate that this parasite has evolved species-specific mechanisms to successfully infect a wide range of hosts.

27 1.2.4.1 Tachyzoite morphology

The crescent shape of the 2-6µm tachyzoite is attributed to its characteristic apical complex that tether phyla-specific organelles, referred to as Dense Granules (GRA),

Micronemes (MIC) and Rhoptries (ROP) assembled in a complex at its anterior end (246).

Hence, the apical end of the parasite contains a vast number of organelles, whereas the nucleus sits centrally (247) and the mitochondria is present in the posterior region (248).

The outer covering of the parasite is known as the pellicle, which consists of an outer and inner membrane (249). The cytoskeletal network provides the parasite’s structural integrity, and its characteristic shape is maintained by subpellicular microtubules (250).

The anterior tip of the apical complex is known as the conoid, which contains two preconoidal rings and subunit structures of unknown composition arranged in a counterclockwise manner to form a tube-like structure (248, 251). Posterior to the conoid, a polar ring functions as a microtubule organizing center. Two microtubules that run from the preconoidal rings and along the length of the parasite are postulated to help direct polarized secretion of effector molecules from the specialized organelles tethered to the apical complex during the invasion process (251). These specialized GRA, MIC and ROP organelles are tightly associated with this microtubule network which facilitates their localization to the conoid region (252). Upon appropriate stimulation, these specialized organelles secrete a large number of effector proteins critical for the invasion process.

Importantly, T. gondii is not motile and thus depends on its cytoskeleton to enable parasite movement across host cell surfaces.

28 1.2.4.2 The active invasion process

The invasion process of host cells involves a highly ordered, concerted chain of events that results in active penetration of the host cell and the formation of a protective vacuole inside of the host cell, which is known as the parasitophorous vacuole (PV)

(Figure 1.3). The parasite begins the active invasion process only after it has become loosely attached to the host cell surface, followed by a gliding movement across the host cell surface before it orients its highly polarized apical end into an active penetration complex. At this point a specialized structure known as the moving junction is established, through which an invagination of the host cell membrane induces the organelles at the apical ends to discharge their contents sequentially. Ultimately this process facilitates entry of the parasite into host cells to establish an intracellular niche inside the PV (253). The invasion process takes less than 90 seconds to complete. The kinetics of protein release from the organelles is associated with numerous specialized functions whereby these organelle-specific effector proteins function to promote the establishment of the intracellular environment that supports parasite growth.

1.2.4.2.1 Cell recognition, attachment and evasion of host defenses

An important prerequisite for infection is the initial interaction with the host cell, which includes adhesion followed by attachment. As a generalist parasite, T. gondii has a unique capacity to invade almost any nucleated cell in a large range of mammals. This suggests that the parasite either binds numerous or common ligands found ubiquitously across all hosts. However, very little is known about specific host ligands that facilitate this process. The interaction of many pathogens with a large number of host cells to initiate infection is typically mediated by proteoglycans (254), which are common carbohydrate

29 structures found on host cells. Previous work has shown that T. gondii possesses a lectin- like activity that is associated with their ability to interact with host sulfated proteoglycans

(SPGs) (255–257). Equally, cells lacking proteoglycans or sulfation have been shown to be more resistant to T. gondii infection (256, 258).

1.2.4.2.1.1 The SRS superfamily of surface coat proteins

The initial contact between parasites and host cells occurs in the extracellular environment. It has been postulated that parasite specific surface coat proteins are responsible for mediating the initial attachment to host cells. The surface of T. gondii is dominated by a superfamily of proteins, known as the SRS (SAG1-related sequence) proteins (259). These proteins are all structurally related to SRS29B (formerly known as

SAG1), and this protein superfamily typically shares from 24-99% sequence identity (260).

SRS proteins are tethered to the surface of the parasite by glycosylphosphatidylinositol

(GPI) anchors (259). Genomic analyses have identified ~180 SRS genes distributed across

T. gondii’s 14 chromosomes at 57 genomic loci (261). An important feature of these proteins is that they are developmentally regulated, with largely non-overlapping expression between parasite stages, suggesting that they have different roles at various stages of the parasite lifecycle. Further, SRS gene family composition is different between strains indicating that different strains possess distinct sets of SRS genes, which may contribute to strain-specific phenotypic differences in attachment, invasion, dissemination and pathogenesis (260).

Early studies showed that antibodies against major surface antigens SRS29B and

SRS34A (formerly SAG2) inhibit host cell infection (262, 263), indicating that SRS proteins are critical for infectivity. Several SRS proteins, including SRS29B and SRS57

30 (formerly SAG3) have been shown to bind SPGs with strong affinity (258, 264). The crystal structure of SRS29B determined the structural basis for these observations, by revealing a dimer dependent basic groove that was well suited to coordinate a negatively charged cellular ligand such as heparan sulfated proteoglycans (265). Based on this structure, the model developed for SRS57 identified a substantially more basic groove consistent with its role as a major surface adhesin capable of recognizing negatively charged SPGs (265).

Although invasion is a relatively quick process (266), the surface coat must directly confront and evade host defenses in order to successfully establish infection. In addition to their role in attachment, SRS proteins have been associated with subversion of host immunity. Expression of SRS29B and SRS34A, two highly immunogenic tachyzoite proteins, is thought to act as immune decoys and attract host immunity against the tachyzoite stage to regulate virulence (260, 267, 268). Several studies have also suggested that SRS29C and SRS29B possess additional immunomodulatory roles and function as regulators of virulence (261) and inflammation (269), respectively, however the precise molecular mechanisms behind these phenotypes remain enigmatic.

Comparative modeling has predicted that the 6-CYS surface proteins encoded by

Plasmodium share structural similarities with the SRS proteins (270). This was confirmed when the crystal structure was solved for Pf12 and revealed that the 6-CYS proteins share tertiary structural homology with the SRS fold (265, 271, 272). Although the precise biological function for the SRS proteins remain debated, two of the 6-CYS proteins are known to promote sexual competency and resist complement activation. Indeed, 6-CYS proteins Pfs47 and Pf92 have been shown to suppress the mosquito complement-like

31 system and to regulate human complement by recruiting the soluble host regulator Factor

H, respectively (168, 171). The evidence for complement evasion mediated by Plasmodium

6-CYS proteins may indicate that the structurally homologous Toxoplasma SRS proteins possess a similar biological role during active infection.

1.2.4.2.2 Actin-based motility

Host cell entry is initiated after host cell surface contact by the parasite’s apical complex; however, this process also requires parasite motility and propulsion to translocate across the host cell membrane. The parasite’s crescent shape and spiraled cytoskeleton contribute to a substrate-dependent corkscrew motility called gliding, which results in a series of circular and helical turns revolving along the longitudinal axis of the cell (273).

This process relies on both actin and myosin present between the plasma membrane and two underlying membranes, referred to as the inner membrane complex. Studies using host cells treated with cytochalasin D, (274) which impair host cell actin filaments, and parasites with mutated actin or myosin genes (275, 276) have shown that T. gondii invasion requires parasite-driven actin-based motility, rather than host actin-driven internalization of the parasite by the host cell. Time lapse video microscopy has established precisely how parasite gliding propels the parasite forward at 1-2 microns per second (273), allowing it to move forward as it enters the host cell and through epithelial layers as it disseminates

(237). The parasite’s ability to disseminate due to its active motility is strain-dependent, where mouse virulent Type I strains display greater migratory capacity compared to avirulent Types II and III. However, recent studies have suggested that Apicomplexan parasite motility may not depend strictly on parasite actin, but may also rely on an endocytic-secretory cycle (277, 278).

32 1.2.4.2.3 The moving junction

The moving junction, a structure conserved among Apicomplexan parasites, is a transient configuration at the site of penetration that allows the parasite to squeeze through the host cell plasma membrane (279, 280). Protein secretion from the parasite’s specialized organelles then aids in the firm attachment and translocation of the parasite, resulting in an invagination of the host cell membrane that ultimately surrounds the parasite and forms the specialized intracellular parasitophorous vacuole. A coordinated discharge of microneme and rhoptry proteins has been associated with this step (253). This process is highly dependent on specific proteins, notably the highly conserved Apicomplexan protein microneme membrane antigen 1 (AMA-1), which complexes with rhoptry neck proteins, known as RONs, to make up the moving junction (281, 282). The moving junction controls internalization by selectively excluding certain host cell plasma membrane proteins based on their membrane anchors (283). For example, GPI-anchored host cell proteins such as

CD55 and Sca-1 are incorporated into the PV whereas CD44 and beta-1 integrin are excluded (283). This selection of host proteins has been postulated to promote a non- fusogenic PV. Remarkably, active entry into host cells is relatively stealth-like, and does not induce host cell membrane ruffling, actin microfilament reorganization, or tyrosine phosphorylation of host proteins (266).

1.2.4.2.4 The parasitophorous vacuole

Internalization of the parasite upon entry into the host cell results in the formation of an intracellular compartment known as the parasitophorous vacuole (PV). A mature PV is a protective, membrane bound, cytoplasmic vesicle that is derived from the host cell

(284) and never fuses with the lysosomal compartment nor acidifies (285, 286), in this way

33 parasites avoid intracellular killing. The parasite resides inside this specialized vacuole in the cytoplasm for its entire intracellular phase, where it rapidly divides. In order to survive intracellularly, the parasite must establish a permissive environment for replication. To do this, the vacuole must exchange nutrients across the PV membrane (PVM). The parasite relies on the host cell for much of its nutrients, as Toxoplasma is an auxotroph for tryptophan, arginine and purine (287–289). This is intriguing considering the PV is totally isolated and is devoid of host cell transporters or receptors of host origin (290). Early studies determined that the PVMs of Apicomplexan parasites are selectively permeable to small molecules (13-19kDa) through pore like structures (291).

The PVM consists of an intravacuolar network (292) that facilitates many of its functions, in particular, the ability to support intracellular parasite survival. It was hypothesized early on that the parasite modifies the PVM to facilitate parasite survival

(290). Secretion of proteins from the dense granules upon invasion and post-invasion can be detected in the vacuolar space within minutes after host cell invasion. Dense granule proteins, known as GRA proteins, have been shown to be directly involved in the formation of the tubules that make up the intravacuolar network (293–295). In recent years, dense granule proteins GRA17 and GRA23 were identified to modify the PVM and facilitate the transport of small molecules (296) across the PVM. To export these effector proteins through the PVM, the parasite encodes proteins ASP5 (297), a Golgi-resident aspartyl protease 5, MYR-1 (298), and MYR3 to form an export complex (299).

1.2.4.2.5 Parasite replication and egress

Toxoplasma replicates inside the PV through a process known as endodyogeny, where two daughter cells are generated within one mother cell (300) every 6-10 hours, until

34 the host cell is lysed (typically) 48 hours post infection, however this varies depending on parasite genotype. At the start of endodyogeny, a scaffold for daughter cells is provided by the inner membrane complex, where two membranes begin to form in the middle of the parasite cell whilst cytoskeletal elements are assembled. Mitochondria, organelles and cytoplasmic content are divided between the two progeny (249, 252, 301). Parasite egress is a rapid, Ca2+-dependent (302) and cytolytic event that results in the release of motile parasites ready to infect new cells. Microneme (303), dense granule, and perforin-like proteins (PLP) (304) have also been implicated in this process of exiting the PVM and host cell.

1.2.4.3 Phagocytic/endocytic uptake

Phagocytic uptake has been reported for Toxoplasma by phagocytic cells, however is not believed to be a major mode of entry for this parasite (266). In contrast to invasion which occurs within 60-90 seconds, Fc-mediated endocytic uptake of Toxoplasma takes 2-

4 minutes and results in extensive membrane ruffling, reorganization of the host cytoskeleton, and phosphorylation of host proteins (266). The endocytosed parasite is then surrounded by a phagocytic vacuole (phagosome) that is distinct from the PV and rapidly matures into a phagolysosome, which acidifies and kills the parasites (305). Parasites coated with nonspecific immunoglobulins have also been shown to enhance phagocytosis but these do not have a large effect on their proliferation (306). Occasionally, parasites internalized by phagocytosis escape from the phagosome (266).

Although active invasion is considered the major mode of entry into the host cell, a notable observation is that parasite entry among parasites deficient in actin or myosin is not completely abrogated (307). Recent work has added to this idea that alternative routes

35 of parasite entry exist, demonstrating a non-canonical pathway of invasion, whereby virulent Type II parasites are first internalized by phagocytosis, followed by active invasion from the phagosomal compartment to form a PV (308). Furthermore, this unique mode of entry was not observed for Type I strains, which were more invasive and it has been suggested that they lack the parasite ligand which is recognized by phagocytic receptors

(308). In this study, the authors hypothesized that this non-canonical pathway results in enhanced immune stimulation and control of acute infection with Type II parasites.

Interestingly, human monocytes infected with T. gondii in vitro resulted in immune stimulation only upon phagocytosis, and not when parasites invaded these cells (309, 310), which is in stark contrast with the murine model. These studies indicate that parasite entry into phagocytes by phagocytic uptake can also play a critical role in influencing immune responses.

1.2.5 The immune response to T. gondii and mechanisms of immune evasion

The emergence of toxoplasmosis as an AIDs defining infection underscored the importance of T. gondii’s interaction with the immune system in determining immune outcome. Both innate and adaptive immune responses contribute to host protection during

T. gondii infection. Detection of the pathogen by non-specific, germline encoded receptors, both membrane-bound and soluble, are critical to the innate immune response, and results in the production of chemokines and cytokines that help shape the adaptive immune response. T. gondii is known to induce a robust inflammatory Th1 immune response that helps eliminate intracellular pathogens. It is ultimately the adaptive immune response that controls long term infection; however, it is incapable of completely clearing the parasite.

Ironically, the immune pressure induced by the adaptive immune response also contributes

36 to stage conversion from the acutely proliferating tachyzoite phase to the chronic persistent bradyzoite phase. Thus, early immune events that precede the development of adaptive immunity are of therapeutic interest.

The long-term co-evolution of the Apicomplexa with their hosts has resulted in sophisticated mechanisms of host evasion that allow parasites to persist (311). T. gondii employs multiple mechanisms to balance immune evasion and immune activation during infection of its intermediate hosts, including the control of host gene transcription and the dysregulation of host immune signaling pathways to alter cell adhesion, migration and secretion of cytokines (312). The pathogenesis of infection is directly dependent on the suite of parasite effector proteins encoded by each parasite strain and how these strain- specific proteins function to subvert host innate immune responses (237). Many of these strategies rely on polymorphic proteins that are secreted from specialized organelles, which include the rhoptry proteins that are directly secreted into the host cell cytosol upon invasion as well as several dense granules that cross the PVM post infection. Often, these effector proteins work in complexes to mediate their regulatory function.

Despite these advances in understanding both host and parasite factors that contribute to immunity and pathogenesis, there also remain several gaps in our understanding of the immune response to T. gondii, which most notably includes the vast differences in immune responses between mice and humans. Likewise, the parasite factors that mediate serum resistance and the evasion of host immunity during its extracellular phase are poorly understood, a point during infection whereby the parasite is not protected by an intracellular vacuole and is most susceptible to immune attack.

37 1.2.5.1 Cellular immunity

1.2.5.1.1 Innate cellular responses

1.2.5.1.1.1 parasite recognition

The innate immune response is responsible for sensing and appropriately priming the adaptive immune response. Th1 cell-mediated immunity is critical for clearing intracellular pathogens, including Toxoplasma (313). This response relies on the production of Th1 cytokines, primarily interlukin-12 (IL-12), interferon-g (IFN-g) and tumor necrosis factor-a (TNF-a). These cytokines drive T cells to directly kill infected cells or help them clear intracellular pathogens by stimulating antimicrobial responses against infected cells. Although stimulation of IL-12 by innate immune cells responding to

T. gondii during acute infection is important for both mice and humans, parasite recognition events between mice and humans that induce Th1 cytokine production are distinct.

In the murine model, it was determined early on that IFN-g was critical to host resistance against T. gondii (314–316), indicating that Th1 cell-mediated immune responses are required for host protection. One of the first cell types to encounter

Toxoplasma after the parasite crosses the intestinal epithelial barrier are antigen presenting cells (APCs). Dendritic cells, macrophages and neutrophils have all been shown to produce

IL-12 in response to T. gondii (317–319). Toll-like receptors (TLRs) are pattern recognition receptors expressed on myeloid cells that play an important role in detecting bacteria, viruses and parasites (320). MyD88 is an adapter protein that facilitates TLR signaling and induces inflammatory cytokine responses. Early studies determined that

TLRs were important in the response mounted against T. gondii because My88 deficient macrophages failed to induce IL-12 production in vitro (318). In vivo studies using MyD88-

38 /- mice have established that signaling through MyD88 is critical for host resistance against

T. gondii, as deficient mice are highly susceptible to infection. This dependence on MyD88 for IL-12 production suggests that parasite sensing by TLR pattern recognition receptors is a critical factor in the induction of Th1 responses. MyD88-dependent activation of IL-

12 was later determined to be dependent on the detection of parasite profilin by TLR11 and

TLR12 (321–323) in distinct subsets of dendritic cells and that this was required for host control of parasite replication during acute infection (324).

However, IL-12 is not the only MyD88-dependent effector that contributes to induction of a protective adaptive immune response, suggesting that there are additional innate stimuli to promote Th1 effector responses in vivo (324). Recently, cytosolic immune sensors, such as NLRP-1 and NLRP-3 (Nod-like receptors), have been shown to induce the inflammasome in response to T. gondii in both murine and human cells (325–328).

Inflammasome activation induces IL-1b (329) and IL-18 production, two cytokines that contribute to Th1 inflammatory responses and the death of infected cells.

In human infection, several studies in patients with acquired toxoplasmosis have supported a role for the Th1 inflammatory response in altering the pathogenesis of T. gondii infection (330). IL-12 is also produced by human monocytes and neutrophils (331) in response to T. gondii. However, TLR11 is a non-functional pseudogene and TLR12 does not exist in humans (311). Thus, humans have distinct mechanisms of parasite recognition that elicit IL-12 production. Unlike murine phagocytes, phagocytosis is thought to be the primary determinant driving IL-12 responses among distinct subsets of monocytes, rather than active invasion (310). This suggests that recognition requires direct contact with live

39 parasites, however, how the human innate immune system detects T. gondii is not well understood.

1.2.5.1.1.2 Strain-specific modulation of innate immune signaling pathways

TLR activation results in the nuclear translocation of transcription factor NF-kB, which regulates the transcription and synthesis of IL-12 and other inflammatory cytokines such as IL-6 and IL-1b (332). Binding of cytokines and interferons can also induce intracellular signaling that results in the activation of STAT transcription factors that can impact IL-12 signaling. These signaling pathways often rely on phosphorylation or activation of domains within the signaling cascade to facilitate transcription factor translocation into the nucleus. Toxoplasma infection has been shown to interfere with both

NF-kB and STAT signaling. In vitro, Type II strains are known to induce much higher levels of IL-12 and IL-1b in comparison to Type I and Type III strains (318, 325, 333).

Early studies demonstrated that infection with Type I strains resulted in rapid STAT3 activation and blockage of NF-kB translocation to the nucleus resulted in suppressed IL-

12 and TNF-a in macrophages infected with Type I strains (334–336). Infection with Type

II, on the contrary, induced p38 MAPK activation and nuclear localization of NF-kB to upregulate IL-12 production (334, 337). Using forward genetic crosses it has been possible to identify several additional parasite effectors that have been shown to interfere with transcriptional regulation and cytokine production in a strain-dependent manner. For example, the parasite effector protein ROP16 was identified as the factor mediating activation of STAT3 and STAT6 in Type I and Type III infected murine macrophages and human foreskin fibroblasts (335, 338). These rhoptry proteins are parasite-specific active kinases that are secreted upon invasion and have been shown to modulate cell effector

40 function, specifically they are known to directly phosphorylate STAT3 in infected host cells (335, 339). The inability of the Type II ROP16 allele to activate STAT6 was later shown to be due to a single polymorphism in the kinase domain of the Type II allele (339).

Thus, these differences observed in vitro suggested that such strain-specific parasite factors regulate cytokine responses in vivo and impact parasite pathogenesis in animal models in a strain-specific manner. Indeed, murine infection with Types I, II and III in vivo have vastly different outcomes in laboratory mice, whereby Type I strains are universally lethal at low inoculums (i.e., 50 tachyzoites) whereas Type II and Type III strains are avirulent, however this varies depending on the mouse genotype and infection inoculum

(236). In vivo studies have established that infection with Drop16 parasites only have a moderate impact on acute virulence in mice (340), indicating that control of host signaling was dependent on additional factors. Additional studies have identified other polymorphic secreted effector proteins that impact parasite pathogenesis. The dense granule protein

GRA15, for example, is a highly polymorphic effector protein that is responsible for activating NF-kB and contributing to higher IL-12 production in Type II infected cells, whereas the Type I allele fails to do so (341). GRA15 has also been implicated in the induction of IL-1b in human monocytes infected with Type II strains (325).

Although the cumulative role of these polymorphic effector proteins initially suppresses innate Th1 responses, they ultimately lead to greater parasite dissemination and higher parasite burden in mice infected with Type I strains in vivo. These mice, ironically, die of an over exuberant, unregulated proinflammatory Th1 response, otherwise known as a cytokine storm (342, 343). It is postulated that early impairment of Th1 responses results in higher parasite burdens and this results in more pronounced tissue damage due to

41 excessive parasite proliferation which ultimately leads to secondary responses and immunopathology (239). Type II strains, however, possess a different combination of polymorphic secreted effector alleles that induce early IL-12 production during acute infection which controls tachyzoite proliferation and fails to induce the cytokine storm that mediates Type I strain pathogenesis (239).

1.2.5.1.1.3 Cell autonomous immunity

IL-12 is important cytokine for driving IFN-g production, which is a key mediator in both mice and humans to control parasite proliferation (315). IFN-g also stimulates cell autonomous immunity in infected cells to help suppress parasite growth and directly kill parasites. IFN-g is produced by NK cells, CD4+ and CD8+ T cells and mediates host cell resistance to T. gondii infection in both hematopoietic and non-hematopoietic cells (344).

IFN-g exerts its effects by inducing immunity-related GTPases (IRGs), guanylate-binding proteins (GBPs), the production of antimicrobial molecules including nitric oxide (NO) and reactive oxygen species (ROS). Specifically, IRGs have been shown to play an intrinsic role in antimicrobial defense in murine cells by targeting, accumulating, and disrupting the

PVM to promote parasite destruction (345). Mice deficient in IRGs are no longer resistant to T. gondii infection (346, 347).

Parasite susceptibility to this response is also strain-dependent, whereas Type I strains are resistant to IRG-mediated effects, Type II and Type III infected cells are susceptible (348). Several parasite effector proteins that interfere with IRGs have been identified. ROP5 is a psuedokinase that stabilizes a complex with ROP17 and ROP18 two kinases that work together to prevent IRG oligomerization by phosphorylating and inactivating murine IRGs to protect the integrity of the PVM in Type I infected murine

42 cells (349, 350). Importantly, neither ROP5 nor ROP18 promote parasite survival in IFN- g activated human cells (351). Although mice encode 23 IRGs and GBPs, humans only encode 2 IRGs, so it was not surprising that ROP5 and ROP18 are dispensable and do not play a role in human defense (352). These studies have demonstrated that mice and humans have vastly different mechanisms to deal with vacuolar intracellular pathogens like T. gondii. Humans possess other IFN-g-inducible genes that are deployed against T. gondii to limit their infection, such as indoleamine 2,3-dioxygenase (IDO1) (353, 354). Although T. gondii is known to interfere with human IFN-g activated genes (355), the parasite factors involved in regulating human cell autonomous immunity are less understood. Several studies have suggested that they are not derived from the rhoptries(356) and have suggested that the dense granule protein GRA15 may be more important in regulating human infection than in mice (325, 354).

1.2.5.1.2 Adaptive cellular-mediated responses

The adaptive immune response differs from innate immunity in that it is highly specific, provides long-lasting protection, and requires more time to develop. These responses are mediated by B and T lymphocytes that rely on complex interactions that include recognition of presented antigen, costimulatory molecules, and signals from innate immune cells in order to differentiate B and T cells into appropriate effector subsets to control Toxoplasma infection. Long term control and prevention of reactivated disease is primarily mediated by T cell mediated immunity, which is comprised of CD4+ and CD8+

T cells. Both subsets of T cells are responsible for resistance to T. gondii infection (357) and for long-term host survival by preventing reactivation of disease (358).

43 Cytotoxic CD8+ T cells mediate protection by producing IFN-g and directly killing

T. gondii infected cells through perforin mediated cytolysis (359). In humans, the emergence of toxoplasmosis is associated with a decline of CD4+ T cell numbers in HIV infected individuals, demonstrating a critical function for CD4+ T cells in controlling reactivated disease. CD4+ T helper cells mediate protection by their production of IFN-g and TNF-a which induces antimicrobial activity in cells infected with T. gondii. In the mouse model, CD4+ T cells have been shown to contribute to optimal B cell (360) and

CD8+ T cell function (361) by upregulating costimulatory molecules and antigen presentation. However, CD4+ T cells and the production of IFN-g also contributes to intestinal immunopathology during acute oral infection in some strains of mice (362). T. gondii is also known to induce immunopathology in the brain during chronic infection

(363). Although both CD4+ and CD8+ T cells are critical for long term survival of mice, both CD4 and CD8 deficient mice do not succumb to acute infection (359). Conversely, the early, acute death of mice deficient in innate immune factors such as MyD88 and IL-

12 demonstrate that stimulation of innate immunity is ultimately critical for resistance to acute infection.

1.2.5.1.3 Humoral immunity

The humoral immune system is comprised of soluble protective factors, which include antibodies and the complement system which are of adaptive and innate origin, respectively. Upon stimulation with antigen, B cell activation leads to the secretion of antigen-specific antibodies. Antibodies mediate their protective effects through several ways: blocking entry into host cells, opsonization of pathogens to target them for

44 phagocytosis, antibody-dependent cell mediated cytotoxicity, or direct lysis via activation of the classical pathway of the complement system.

Although infection with T. gondii induces a robust antibody response (364)(365), humoral immunity initially did not appear to play a significant role in mediating protection during acute infection. Early studies revealed that serum transferred from infected hamster to naïve hamsters did not provide significant protection against T. gondii challenge (366).

Studies in B cell deficient mice, however, showed that these mice were less resistant to T. gondii infection than wild type mice, despite no real changes in IFN-g, TNF-a, and NO synthase (367). Because B cell deficient mice succumbed to infection 3-4 weeks after infection, this suggested that antibodies were dispensable during acute infection; however, these mice had higher cyst burdens and more pathology in the brain indicting that antibodies do play an important role in controlling chronic infection.

During early acute infection, the early humoral response is restricted to just two antigens, the T. gondii surface antigens SRS29C and SRS34A (368). It is postulated that these antigens help to control parasite proliferation while also inducing stage conversion.

Interestingly, intraperitoneal injection of monoclonal antibodies against two T. gondii surface antigens (35kDa and 14kDa) conferred total protection when challenged with a moderately virulent strain (369). Antibodies also provide protection by activating the classical pathway of the complement system, as T. gondii specific antibodies have been shown to kill live tachyzoites and this is the basis for the Sabin-Feldman dye test (370,

371), which is commonly used to diagnose T. gondii infection.

Arguably, one of the least characterized immune responses to T. gondii is the complement response during acute infection in non-immune hosts. To date, only one study

45 has investigated the complement response to T. gondii in non-immune human serum (173).

In this study by Fuhrman and Joiner, T. gondii was shown to primarily activate the alternative pathway and parasite resistance to complement-mediated lysis was achieved by the inactivation of C3b. Several parasite surface proteins, including p30 and p22, were shown to be modified by the complement effector protein C3b (173). p22 and p30 are predicted to be SRS29B and SRS34A, however the identities of these proteins were never resolved. Moreover, the molecular basis for complement inactivation and the contribution of serum resistance to pathogenesis remain poorly understood. Considering the contribution of parasite recognition by innate immunity to host resistance against T. gondii, the diverse roles of complement system in pathogen detection, and its ability to bridge adaptive immune responses, this thesis set out to understand how the complement system impacts T. gondii acute infection.

1.2.6 Hypothesis, Specific Aims, and Experimental Approach

The goal of this thesis work was to determine the parasite surface factors regulating complement activation and to determine the mechanism of C3b inactivation. We hypothesized that T. gondii SRS proteins are important factors in this interaction and that this interaction with the complement system contributes to their biological function.

In Research Aim 1, we developed flow cytometry-based tools to compare complement activation and serum resistance in Type I and Type II strains incubated in the presence of non-immune human serum. We identified strain-specific differences in C3b deposition between Type I and Type II strains. We then took a forward genetic approach by utilizing recombinant progeny from an experimental cross between a Type I x Type II strain to map parasite genes responsible for the observed strain-specific difference in C3b

46 deposition. Using this unbiased approach, we identified two surface coat SRS proteins,

SRS29C and SRS25, that exhibited distinct yet cooperative mechanisms of lectin and alternative pathway regulation by recruiting host regulators Factor H and C4b-binding protein. These findings are described in Chapter 2.

In Research Aim 2, we took a structural approach to investigate the precise molecular basis for SRS29C-mediated Factor H recruitment. In Chapter 3, we solved the crystal structure of SRS29C and showed that it possesses a dimer dependent basic groove that is capable of docking host heparan sulfated proteoglycans, a “self” host factor recognized by host regulator Factor H. We then used biochemical analyses to confirm

SRS29C heparan binding and engineered a SRS29C transgenic over expressing strain with mutations in key lysine residues critical for binding SPGs to show that SRS29C proteins that fail to bind SPGs abrogate Factor H recruitment. The biological relevance of these interactions was tested in vivo using transgenic SRS29C over expressing strains, SRS29C deficient parasite strains, and C3 deficient mice. We showed that SRS29C expression regulates pathogenesis and alters the Th1 cytokine production in an SPG- and C3- dependent manner in vivo.

In Research Aim 3, we investigated the capacity of SRS57 to bind host SPGs as had been previously predicted by comparative modeling studies and whether SRS57 was capable of regulating complement deposition on the parasite surface. In Chapter 4, we engineered SRS57 deficient parasites to evaluate SRS57 interactions with both C3b and

FH and showed that SRS57 is a C3b acceptor capable of binding complement proteins. We also generated a de novo crystal structure for SRS57 and showed that it neither dimerized nor supported a basic groove necessary to coordinate SPGs. In vivo characterization of

47 SRS57 showed that SRS57 is capable of regulating the pathogenesis of T. gondii infection in a C3-dependent manner by controlling parasite load.

48 1.2.7 Figures

CLASSICAL LECTIN ALTERNATIVE PATHWAY PATHWAY PATHWAY

C1s MASP-1 C3

MASP-3 C1r H2O

C1q MBL/ficolin C2 C3(H2O) C4 Factor B Factor D

monocyte C3 AP amplification loop C3 CONVERTASE B cell mast cell Neutrophil CR2 C3a C3b

antigen + C3aR C3dg C5aR Opsonization

C5 CONVERTASE INFLAMMATION C5b CR3 CR4 C5a C5b-9 Phagocytosis

Cell Lysis

Figure 1.1. The complement cascade and effector functions. The complement system is activated via three separate pathways that have distinct recognition mechanisms. Complement activation results in the formation of pathway-specific C3 convertases. All three pathways converge at C3 activation, which generates effector molecules C3a and C3b. C3a is recognized by cognate receptor C3aR to mediate recruitment of innate immune cells and promote inflammation. C3b is covalently coupled to pathogen surfaces and promotes opsonization. C3b is recognized by complement receptors CR3 and CR4 to promote phagocytosis and by CR2 on B cells to stimulate adaptive immunity. C3b allows for progression to the terminal pathway by forming C5 convertases. The terminal pathway requires C5 activation product C5b to initiate the assembly of the membrane attack complex (MAC, C5b-9). The pore inserts into the cell membrane and promotes cell lysis. This figure was generated using BioRender.

49 Table 1.1. Host regulators of the complement system and their targets.

Both soluble and membrane bound regulators target several points of the complement cascade, including C3/C5 convertases, C3b, C4b, and C5b-9 pore formation.

50 Table 1.2. Intracellular protozoan parasite evasion of the complement system.

Trypanosoma cruzi, Leishmania, and Plasmodium parasites encode parasite-specific complement regulators that mimic the function of host regulators, or recruit host regulator proteins to down regulate complement activation.

Figure 1.2. T. gondii life cycle. The Toxoplasma life cycle generates two infectious forms: sexual stage oocysts containing sporozoites and asexual stage tissue cysts containing bradyzoites. The ingestion of vegetation or water contaminated with oocytes or meat containing tissue cysts from an infected animal ensures transmission to both herbivores and carnivores. Upon ingestion, excystation of parasites in the gut of the intermediate host results in the differentiation into the rapidly replicating asexual form, the tachyzoite. The tachyzoite disseminates from the gut throughout the host and is able to cross various biological barriers, including the placental and blood-brain barrier. The immune pressure of a strong Th1 response clears the tachyzoites but also drives differentiation into the chronic stage, resulting in the formation of tissue cyst containing slowly replicating bradyzoites. Should the definitive host prey on infected animals, this results in the sexual cycle, where differentiation into male and female gametes in the feline gut results in the formation of oocysts that are shed into the environment. This figure was created using BioRender.

Figure 1.3. T. gondii invasion. The invasion process involves several stages: (1) initial attachment to the host cell surface mediated by SRS (SAG1-related sequences) proteins, (2) gliding across the host cell surface to orient apical end, (3) firm attachment, secretion of microneme proteins, and formation of the moving junction, (4) active penetration into the host cell and secretion of rhoptry proteins into the host cell cytosol, and (5) the formation of the parasitophorous vacuole and establishment of an intracellular niche

Figure adapted from: Alexander DL, Mital J, Ward GE, Bradley P, et al. (2005) PLoS Pathog 1(2)

53 CHAPTER 2: SERUM RESISTANCE IN TOXOPLASMA GONDII IS

REGULATED BY SURFACE COAT PROTEINS SRS25 AND SRS29C1,2

2.1 Introduction

At early stages of infection, intracellular protozoan parasites are vulnerable to humoral attack in the extracellular environment. The complement system, a humoral effector system that activates on pathogen surfaces, is considered to be the first line of defense. Complement activation is initiated by three pathways, known as the classical, lectin, and alternative pathways. Recognition of immune complexes and microbial carbohydrates activate the classical (CP) and lectin (LP) pathways, respectively. The alternative pathway (AP) activates on surfaces that cannot control the spontaneous hydrolysis of C3, which are typically non-self surfaces that lack host regulator proteins.

Recognition triggers a sequential cascade of proteolytic events that results in the assembly of complexes known as convertases, that generate active proteins C3a and C3b. These proteins mediate the biological effects of the complement system, which include forming a pore complex to mediate direct lysis, promoting inflammation, targeting pathogens for phagocytosis, and stimulating adaptive immunity (63). However, excessive complement activation can also result in damage and inflammation in nearby tissues, thus it must be tightly regulated (372).

______1Citation: Sikorski, P.M., Commadaro A.G., and Grigg M.E. Toxoplasma gondii recruits C4b-binding proteins and Factor H to mediate serum resistance and promote parasite persistence in vivo. Under review Frontiers in Immunology. 2Citation: Sikorski, P.M., Pszenny, V., Sardinha da Silva, A., Khan, A., Gray, T., Commodaro, A.G., Kennard, A., Boulanger M.J., and Grigg M.E. Toxoplasma surface coat protein SRS29C recruitment of Factor H to evade complement and regulate parasite virulence is sulfated proteoglycan dependent. In preparation.

54 Several intracellular protozoan parasites including Toxoplasma, Leishmania,

Plasmodium, and Trypanosoma, have evolved efficient subversion strategies to evade complement function. Each of these parasites target C3 convertases and the activation product C3b to impede direct killing (165, 167, 172, 173, 373–375). Several surface- expressed parasite molecules that functionally mimic or sequester host complement regulatory proteins have been identified in trypanpsomastids (gp160, LPG, gp63) and

Plasmodium spp. (Psf47, Pf92) (159, 166, 168, 171). To date, however, only one study has shown that T. gondii is resistant to complement killing in non-immune serum by inactivating C3b. The authors concluded that two parasite surface proteins, p22 and p30, were covalently modified by complement activation products (173). However, the identity of those proteins and the mechanism of C3b inactivation remains enigmatic.

T. gondii is a highly prevalent protozoan parasite that can infect essentially any cell in all mammals, including humans. Toxoplasma is comprised of several genotypically variant strains that have been shown to differ in their virulence across a wide range of hosts

(228, 232, 238, 376–381). Type II strains are most prevalent in human and animal infections in North America and Europe (228, 229, 243). Less frequently, human infection with Type I strains or atypical strains with Type I alleles have been associated with causing encephalitis in AIDS patients (241) or recurrent ocular disease in otherwise healthy people

(244). In order to establish infection and cause disease in a large number of hosts, T. gondii employs large families of polymorphic effector proteins to modulate host immune responses. Murine studies have identified several polymorphic secreted effector proteins, including rhoptry and dense granule proteins, that manipulate intracellular immune signaling (335, 338, 382). However, parasite factors orchestrating resistance to host

55 defenses during the parasite’s extracellular phase, including the complement system, are still poorly characterized. Previous work identified two Toxoplasma surface associated proteins, p22 and p30, that resolve by SDS-PAGE at 22 and 30 kDa respectively, that were covalently modified by C3b (173), but their identify was not resolved.

The surface coat of T. gondii is dominated by a developmentally-expressed superfamily of ~180 proteins known as the SAG1-Related Sequences, or SRS proteins

(383). This superfamily of virulence factors and adhesins are thought to function in a biologically quasi-redundant manner, and specifically facilitate interactions with host cells and/or activate host immune responses (260, 265). Several antibody-blocking studies suggested that SRS proteins SRS29B (formerly SAG1) and SRS57 (formerly SAG3) mediate attachment to host cells (258, 384, 385). The crystal structure of SRS29B identified a ligand binding domain that formed when it homodimerizes, replete with a positively charged groove that is capable of docking anionic heparan sulfated proteoglycans (265), consistent with an adhesive property.

The SRS gene family is highly polymorphic and exhibits strain-specific expression across T. gondii genotypes (260, 261). A previous study showed that one SRS protein,

SRS29C (formerly SRS2), is weakly expressed in mouse-virulent Type I strains but highly expressed in mouse–avirulent Type II strains (261). This study also showed that transgenic overexpression of SRS29C in a Type I strain was sufficient to alter its virulence kinetics, suggesting that SRS29C is a negative regulator of parasite virulence during acute murine infection (261). However, the precise biological function for the SRS proteins remains unclear. Previous comparative modeling work suggested that the 6-CYS superfamily of 14 proteins encoded by Plasmodium share a structural fold with the SRS proteins (270). This

56 was later confirmed when the crystal structure was solved for the Plasmodium 6-CYS protein Pf12, revealing a tertiary structural homology with the SRS domain fold (265, 271,

272). The 6-CYS proteins have been shown to promote sexual competency and resist complement activation, and two 6-CYS proteins, Pfs47 and Pf92, have specifically been shown to suppress the mosquito complement-like system or regulate human complement by recruiting the soluble host regulator Factor H, respectively (168, 171). This evidence for complement evasion mediated by the Plasmodium 6-CYS proteins establish a rationale to investigate whether the structurally homologous Toxoplasma SRS proteins likewise regulate the complement system during active infection.

In this study, we hypothesized that SRS surface proteins play an important role in regulating the complement system. To answer this question, we developed a flow cytometry-based assay to measure C3b deposition on the parasite surface. Using this assay, strain specific differences in C3b deposition between T. gondii Type I and Type II strains were observed. We took advantage of this difference to perform a non-assumptive forward genetics screen using recombinant progeny from a I x II genetic cross to map the parasite factors controlling this phenotype. Two SRS proteins, SRS29C and SRS25 were identified.

Using reverse genetics, we determined these SRS proteins exhibited distinct mechanisms for regulating C3b deposition by interacting with host regulatory proteins Factor H and

C4b-binding protein. The relative contributions of C4BP and FH binding to SRS29C and

SRS25 revealed distinct yet cooperative roles for lectin and alternative pathways in response to T. gondii in non-immune human serum.

57 2.2 Results

2.2.1 Parasite genotype affects C3b deposition but not C3b inactivation or serum resistance

Previous work showed that the surface coat of Toxoplasma Type I and Type II strains was covalently modified by C3b (173).To confirm this finding, several assays were developed to measure complement activation and resistance. Previously, radiolabeled C3 was added to non-immune human serum (NHS) to detect levels of C3b that bound the parasite surface upon activation (173). C3 is the most abundant complement protein in human serum (~1.0 mg/ml), thus rapid saturation on the parasite surface may impact the sensitivity of measuring C3b deposition. Therefore, we developed a flow cytometry-based assay using monoclonal antibodies specific for human C3b and incubated parasites in serially diluted serum (2-fold dilutions from 40% NHS-2.5% NHS) to determine the optimal concentration that gave good sensitivity without rapid saturation. Whereas 20% and 40% NHS rapidly saturated within minutes, and 5% and 2.5% were less sensitive in the absolute level of C3b detected, we found that 10% NHS was the optimal concentration, it did not saturate in minutes and was still sensitive enough to detect variable levels of C3b deposited depending on the conditions used. (Supplemental Figure 2.1).

The level of C3b deposition using 10% NHS was determined for two T. gondii strains, Type I strain RH and Type II strain ME49. Contrary to previous work (173), our method identified that Type II strains are coated in a time-dependent manner with significantly higher levels of C3b than Type I strains (Figure 2.1A, 2.1B). Increased deposition of C3b on the surface of Type II strains was a general trait, as equivalent levels and kinetics of C3b were found on Type II strains derived from different animal hosts

58 (ME49, California sheep) and different geographies (CZ, Czech Republic tiger) (Figure

2.1C).

One limitation of this flow cytometric assay is that it cannot distinguish between active versus inactive forms of C3b. Western blot studies were thus performed to assess the form of C3b covalently associated with parasite surfaces and showed that C3b is inactivated within 10 minutes (Figure 2.1D). The degradation pattern of C3b was consistent with host mediated inactivation by co-factor Factor H and serine protease Factor

I (active C3b a 110 kDa, b 75 kDa; inactive iC3b 68 & 43kDa, C3dg 41kD). Although there are significant differences in the quantity of C3b between Type I and Type II strains, the western blot analysis showed that there is no absolute difference in the inactivation of

C3b between strain types.

To determine whether the inactivation of C3b was sufficient to obviate attack complex formation (C5b-9) to prevent serum killing, parasites viability was evaluated using increasing amounts of NHS. Previous work measured viability of parasites exposed to serum using a relatively insensitive trypan blue exclusion test (173). Here, we utilized a fixable amine-reactive viability dye to measure resistance to serum killing by flow cytometry as well as a monoclonal anti-human C5b-9 (membrane attack complex) antibody to measure viability and pore formation, respectively. Despite differences in the absolute levels of C3b, Type I and Type II strains remained equally viable in the presence of non- immune serum, independent of serum concentration. Type II parasites had higher levels of

C3b deposited on their surfaces when incubated in non-immune serum, however, they were still resistant to C5b-9 pore formation, (Figure 2.1F.) suggesting that the mechanism of

C3b inactivation was not strain dependent. As a positive control for complement meditated

59 lysis, parasites were incubated in the presence of immune serum, in which parasites are susceptible to classical pathway activation (371), and both strains were equally susceptible to lysis in immune serum (Figure 2.1E).

Taken together, our data demonstrate that inactivation of C3b, regardless of the levels deposited, was sufficient to prevent serum killing by both Type I and Type II strains.

Moreover, the results suggest that the mechanism of inactivation is the same for both strains. Given the significant differences in C3b deposition between the two strains, the parasite factors regulating C3b levels could potentially be mapped using a forward genetic approach.

2.2.2 Loci associated with regulating C3b deposition are identified by forward genetics

To determine the genetic basis regulating strain-specific deposition of C3b we tested two cat competent, drug resistant strains previously used for a I x II cross, a sinefungin-resistant Type I GT1 strain (GT1-SNFR) and a 5-fluoro-2’-deoxyuridine- resistant Type II ME49 strain (ME49-FUDRR) (382) to formally establish a difference in

C3b deposition between the two parental strains (Supplemental Figure 2.2). A 2-3 fold difference in C3b deposition was identified between GT1-SYNR and ME49-FUDRR strains which was sufficient to perform a forward genetic screen. C3b deposited on the surface of

I x II recombinant progeny was determined for each of the 23 strains from the I x II genetic cross (Figure 2.2A). Two independent experiments were performed in triplicate and the majority of progeny had similar results between experiments. The presence of an intermediate C3b binding phenotype among progeny that did not segregate equally to either parent suggested that the phenotype was multi-genic, rather than controlled by a single

60 locus (Figure 2.2A). Thus, mean MFI’s were used to map this quantitative phenotypic trait.

A previous genetic map of 1,063 single nucleotide polymorphisms distributed throughout the T. gondii nuclear genome (representing 92% coverage) was used successfully to identify the ROP5 virulence locus (382). Here, QTL mapping using R/qtl software revealed three QTL peaks with logarithm of odds (LOD) scores above 2.0, on chromosomes V (222 genes), VIII (337 genes) and IX (101 genes) (Figure 2.2B).Although high density SNP maps have been shown to improve QTL mapping, they can also hamper the LOD score (386, 387). The low LOD scores identified in our analysis may reflect the high resolution map or the complexity of the trait, and thus may have negatively impacted the statistical calculations (388) typically designed to detect single QTLs. It is possible that using less markers or re-sampling would improve the statistical power, however, the QTL peaks were exquisitely distinct with little to no background throughout and each peak was in a region of high marker density, suggesting a complex trait with multiple genes.

To prioritize genes identified within the QTLs responsible for this phenotype, the following criteria were employed to generate a list of candidate genes for reverse genetic analyses: (1) the presence of a predicted signal peptide, (2) gene expression in the tachyzoite stage (261, 389) (toxodb.org), (3) a parasite encoded protease, or 4) a surface coat SRS protein. (Supplemental Figure 2.3A). Using these criteria, we identified several potential candidates, including multiple SRS proteins and two proteases. Other protozoan pathogens, for example Leishmania, encode proteases to degrade complement (163), thus two proteases, TGME49_274110 (Chrm VIII) and TGME49_290840 (Chrm IX) with predicted metalloendopeptidase and serine protease activities, respectively, were

61 considered. The Plasmodium surface coat 6-CYS protein Pf92 is structurally homologous to the Toxoplasma SRS proteins and has been shown previously to recruit host regulator proteins to inactivate C3b, so all 7 SRS proteins located within the top 3 peaks were prioritized for deletion.

SRS proteins were further prioritized based on their expression in the tachyzoite stage (261, 389) and those that had strain-specific differences in their expression. Of the

SRSs identified, only SRS29C on Chrm VIII had a strain-specific difference in its level of expression in the tachyzoite stage. SRS29C expression level differences between Type I and Type II strains (Supplemental Figure 2.3B) also correlated with C3b deposition, so

SRS29C was prioritized for reverse genetic validation. SRS25 on Chrm V was also highly expressed, but no data existed for its strain-specific expression (261). Notably, it is unique in Toxoplasma; it is the only SRS protein that is not developmentally regulated, so it is expressed in all stages of the parasite lifecycle, indicating that it could play an important role in complement regulation for both intermediate and definitive hosts. Due to these restrictive selection criteria, SRS29C and SRS25 were prioritized for reverse genetic studies to test whether they regulate C3b deposition on the surface of Toxoplasma tachyzoites.

2.2.3 Generation of SRS29C and SRS25 knockout strains

To validate the QTL analysis, Δsrs29C and Δsrs25 strain were generated in the

Type II CZ1 Δku80 Δhxgprt and the Type I RH Δku80 Δhxgprt parental strains. The

HXGPRT positive and negative selectable mini-gene was inserted into either the SRS29C or SRS25 locus of Toxoplasma by double crossover homologous recombination using a

CRISPR/Cas9-based gene editing strategy. A single guide RNA (sgRNA) was designed for inserting the HXGPRT gene within the 5’ end of the SRS29C or SRS25 locus

62 immediately after the predicted signal peptide. The HXGPRT gene cassettes was flanked by a 40bp sequence that possessed homology with SRS29C or SRS25 immediately upstream and downstream of the Cas9 cut site to facilitate repair of the DNA double-strand break by homologous recombination (Supplemental Figure 2.4A). Following selection with mycophenolic acid and xanthine for parasites with HXGPRT inserted, clones were isolated and PCR was used to check for the specific insertion of the HXGPRT gene using primers flanking the 5’ end of the SRS gene and a reverse primer inside the HXGPRT gene.

The predicted size of the insert inside the SRS29C and SRS25 loci was 2,430bp and

1,911bp, respectively (Supplemental Figure 2.4B). SRS29C deletion was further verified by staining for SRS29C surface expression using flow cytometry (Supplemental Figure

2.4C).

2.2.4 SRS29C promotes C3b deposition as a C3 acceptor

Because higher SRS29C expression in Type II strains correlated with higher levels of C3b deposition, it was hypothesized that Δsrs29C parasites would show a corresponding reduction in C3b deposition compared to the parental Type II strain. Accordingly, Type II

Δsrs29C parasites exhibited significantly lower C3b deposition (Figures 2.3A, 2.3B).

However, this reduction in C3b deposition was not complete, further supporting the QTL mapping that indicated several parasite derived factors regulate the level of C3b deposited on the parasite surface. To test the sufficiency of this finding, a Type II allele of SRS29C that had been transgenically overexpressed in the Type I strain SRS29C (RH SRS29C++), which only weakly expresses SRS29C (261), was found to possess significantly increased

C3b deposition over the WT RH strain (Figures 2.3C, 2.3D).

63 Because C3b binding correlated with SRS29C expression, these data indicated that

SRS29C was likely a nucleophilic target for C3b. C3 activation results in the exposure of an internal thioester that reacts with nearby nucleophiles, known as acceptors, to form a covalent complex with its target (42, 43, 62). Western blot analysis showed that SRS29C formed a covalent complex with a heat labile serum component in the presence of 10%

NHS, but not with heat inactivated serum (Figure 2.3E).

To determine which SRS proteins were modified by C3b, Toxoplasma surface coat proteins from parasites treated with 10% NHS were immunoprecipitated using an anti-C3 antibody. Prior to Western blotting, eluted proteins that bound to the C3-specific antibody were incubated in the presence of GPI-PLC, which exposes a cross-reactive determinant

(CRD) that is present in the GPI anchor of all Toxoplasma SRS proteins and is detectable by Western blot. Eluted complexes were also treated with 1M hydroxylamine to break the covalent interaction. Probing the treated eluates with an anti-CRD antibody allowed for the detection of all SRS proteins that were covalently modified with C3 and identified a single band at 36kDa that was consistent with the size of SRS29C. (Figure 2.3F). These results indicated that the decrease in C3b deposition on the surface of Δsrs29C parasites was due to a loss of SRS29C as a C3b acceptor site on the tachyzoite surface. While the absence of

SRS29C resulted in a decrease in C3b deposition, significant levels of C3b were still observed on the surface of the parasites, indicating at least one other factor that was not resolved using this analysis.

2.2.5 SRS25 limits C3b deposition

To evaluate whether SRS25 likewise influences C3b deposition, we engineered an

SRS25 deficient strain in both the Type I RH and Type II CZ1 background. Surprisingly,

64 Dsrs25 parasites exhibited significantly higher levels of C3b deposition (Figure 2.4A –

2.4C). One possible explanation for increased C3b deposition is a compensatory over expression of SRS29C, which has been previously shown to occur when two SRS proteins were deleted, specifically in the Dsrs29B Dsrs34A DKO strain (261). Thus, the expression of SRS29C and several other abundantly expressed tachyzoite SRS proteins were measured for their expression level by flow cytometry in the Δsrs25. Although a modest reduction of expression was found for three of the four abundantly expressed surface antigens, SRS29B,

SRS34A, and SRS29C there was no change observed for SRS57 (Supplemental Figure

2.5A).

Another possibility for increased C3b deposition in the absence of SRS25 is the loss of an interacting parasite or host complement regulator, thereby resulting in the unregulated C3b accumulation on the parasite surface. To test this hypothesis, levels of

C3b catabolites iC3b and C3dg on the parasite surface were assessed by western blot. In accordance with flow cytometry results that showed similar levels of C3b on both the

RHDsrs25 and WT RH at 20 minutes, western blot analysis showed that both strains exhibited similar levels of C3b alpha and beta chains (110kDa, 75 kDa) (Figure 2.4C,

2.4D). However, there was a reduction in the C3b inactivation product C3dg (arrow,

39kDa) in the absence of SRS25 (Figure 2.4D). This result supports the possibility that

SRS25 was indirectly involved in limiting C3b levels by facilitating its inactivation by a host regulator or parasite protease. Overall, the data indicated that SRS29C and SRS25 are involved in regulating levels of C3b, but that their mechanisms of doing so are distinct.

65 2.2.6 Recruitment of soluble lectin and alternative pathway regulators C4BP and FH to the Toxoplasma surface reveals a role for tachyzoite surface carbohydrates in lectin pathway activation

Two soluble host proteins that regulate complement activation in non-immune serum are lectin pathway (LP) regulator C4b-binding protein (C4BP) and alternative pathway (AP) regulator Factor H (FH). C4BP primarily targets LP C3 convertases to prevent further cleavage of C3 into its activation products C3a and C3b, however it also is a co-factor for Factor I mediated cleavage of C4b and C3b (108, 390). FH regulates AP activation by facilitating the decay of AP C3 convertases but also acts a co-factor for Factor

I mediated inactivation of C3b into iC3b and C3dg (391). Previous studies have shown that

P. falciparum 6-CYS protein Pf92 regulates complement by recruiting AP regulator FH

(168). To investigate whether Toxoplasma utilizes SRS proteins, which are structural homologs of the 6-CYS proteins, to down regulate complement activation, the binding of host soluble regulators were evaluated by western blot.

Parasites incubated in 10% NHS recruited both C4BP and FH to the parasite surface as demonstrated by Western blot analyses (Figures 2.5A, 2.5B). These data suggested that

Toxoplasma specifically recruits both regulators of the lectin and alternative pathways.

Evaluating the relative contribution of both LP and AP to C3b deposition could provide some insight into the activating nature of Toxoplasma surface coat proteins. To determine the contributions of LP and AP activation to C3b deposition in 10% NHS, the Ca2+- dependent LP was inactivated with 10mM MgEGTA, and thus any C3b deposition measured on Type II ME49 parasites was due to AP activation. Contrary to previous findings (173), these results showed AP activation contributed little to C3b deposition,

66 indicating that LP activation was the major contributor to C3 deposition (Supplemental

Figures 2.6A, 2.6B).

To confirm the role of the LP in C3b activation on the tachyzoite surface, human serum from an individual with a mutation in the mannose binding lectin (MBL) gene was tested. This mutation prevents MBL oligomerization required for carbohydrate recognition, and subsequently impairs LP-mediated complement activation (392). When

ME49 parasites were incubated in the MBL deficient serum, a reduction in C3b deposition was observed compared to normal human serum with functional MBL (Supplemental

Figure 2.6C, 2.6D). This reduction was not as complete as MgEGTA treatment; however, this was expected due to the presence of homologous carbohydrate recognizing molecules such as ficolins, that also activate the LP. LP activation suggests that Toxoplasma harbors surface carbohydrates that can be specifically recognized by the LP pathway.

LP recognition of carbohydrates on microbial surfaces is mediated by two families of lectin proteins, a subfamily of C-type lectins which include mannose binding lectin

(MBL), and ficolins. MBL recognizes terminal monosaccharides such as glucose, mannose, or N-acetylglucosamine (GlcNAc) in a Ca2+ dependent manner (393, 394).

Ficolins are similar in structure and bind acetylated compounds such as GlcNAc and N- acetylgalactosamine (GalNAc) (395). Because little is known about the composition of carbohydrates on the surface of Toxoplasma, a panel of lectins that react with carbohydrates that activate the LP was assessed by flow cytometry. A strain-specific difference in the binding of lectins specific for mannose, GlcNAc, and galactose residues, but not for GalNAc was identified, whereby Type II strains bound more lectin than Type I strains (Supplemental Figure 2.7A, 2.7B). The difference in lectin binding also correlated

67 with levels of C3b activation between Type I and Type II strains. To evaluate whether differences in parasite surface carbohydrates influenced LP recognition, binding of MBL on Type I (RH) and Type II strains (ME49) was measured by flow cytometry. Accordingly, the increased levels of carbohydrates observed on Type II strains correlated with higher levels of MBL compared with Type I strains (Supplemental Figure 2.7C).

SRS29C is glycosylated and resolves as 4 electrophoretic variants by SDS-PAGE

(383). Correspondingly, Δsrs29C parasites exhibited reduced lectin binding

(Supplemental Figure 2.7D), whereas no change was observed for ∆srs29B parasites

(Supplemental Figure 2.7E). To test if glycans bound to SRS29C affected MBL recognition, MBL binding was measured by flow cytometry. In the absence of SRS29C, significantly less MBL binding was observed (Supplemental Figure 2.7F), suggesting that these parasites lack carbohydrates capable of LP recognition. A complete reduction in lectin binding was not observed in the absence of SRS29C, however, suggesting that additional glycosylated parasite surface factors may contribute to complement activation.

Taken together, our data suggest that SRS29C glycans activate the LP pathway and that strain-dependent MBL recognition is the result of different expression levels of

SRS29C between Type I and II strains. To further understand the degree to which the LP and AP are regulated by C4BP and FH, characterization of their contributions to serum resistance would shed greater light onto which pathway is a greater target for T. gondii.

2.2.7 Rapid death of Δsrs25 Δsrs29C DKO parasites establishes that FH and C4BP recruitment determine serum resistance

Because SRS29C and SRS25 impact the level and inactivation of C3b, the capacity for these proteins to bind host regulators C4BP and FH was assessed by flow cytometry.

68 Interestingly, both SRS proteins interacted with C4BP and FH, but to varying degrees

(Figures 2.6A, 2.66B). C4BP binding was abolished in the absence of SRS29C, but only partially reduced in the absence of SRS25 (Figure 2.6A). No reduction in C4BP was observed in the absence of SRS29B, which served as a necessary specificity control for a general loss of surface coat antigen density. In contrast, FH binding was abolished in the absence of SRS25, but only partially in the absence of SRS29C indicating that SRS25 is primarily responsible for FH recruitment. (Figure 2.6B). Importantly, in the absence of both SRS25 and SRS29C, recruitment of both C4BP and FH was abolished in

Δsrs25Δsrs29C DKO parasites (Figure 2.6A, 2.6B). These findings suggest that SRS29C is primarily responsible for C4BP recruitment and supports a dual role for SRS29C in promoting both C3b deposition as an acceptor molecule, while simultaneously recruiting host regulators to facilitate its inactivation.

Conversely, SRS25 critically mediates FH recruitment to regulate C3b deposition.

The reduced levels of iC3b on the surface of Δsrs25 parasites shown in Figure 2.4D is consistent with the loss of FH demonstrated in Figure 2.6B. These results explain the significant increase in C3b deposition in the absence of SRS25 (Figure 2.4A-2.4C).

Moreover, the Δsrs25Δsrs29C DKO parasites phenocopied the ∆srs25 parasites with increased levels of C3b deposition and a similar reduction in the expression of the abundantly expressed SRS proteins SRS29B, SR34A, and SRS57 (Supplemental Figure

2.8A, 2.8B).

To gain further insight into the mechanism of serum resistance, we exploited the differential binding capacities of SRS29C and SRS25 for C4BP and FH, respectively, to deduce the relative contributions of these regulators to parasite resistance. Thus, a serum

69 killing assay was performed to evaluate parasite survival in non-immune human serum

(NHS) (Figure 2.6D). Remarkably, over 95% of the Dsrs25 Dsrs29C DKO parasites were susceptible to complement-mediated killing within 5 minutes of exposure to as little as

10% NHS (Figure 2.6D). Dsrs25 deficient parasites were equally susceptible to complement mediated killing in 10% NHS, but it took longer (~30 minutes) to achieve the same level of serum killing as the DKO parasites (Figure 2.6D), whereas no loss in viability was observed for Dsrs29C parasites (Figure 2.6D) indicating that recruitment of

FH is more important than C4BP to protect parasites from serum killing. Associated with serum killing was a corresponding increase in the levels of C5b-9 MAC complex formation observed for both the Dsrs25 and Dsrs25Dsrs29C DKO parasites (Figure 2.6C) which were capable of proceeding to the terminal pathway whereas this did not occur for Dsrs29C parasites. These data together suggest that C4BP was dispensable for parasite protection and FH was responsible for the majority of serum resistance.

FH is foremost a regulator of the AP amplification loop by its ability to inactivate

C3b (52). Several studies have demonstrated that AP utilizes C3b deposited by both the classical and lectin pathways as platforms to generate an amplification loop to produce more AP convertase and ramp up C3 activation (54, 57). Therefore, these data indicate that the AP mediated amplification of the complement response may be a major target of T. gondii to promote serum resistance in non-immune human serum.

2.2.8 Factor H enhances serum resistance by regulating the amplification of the complement response

To test whether Factor H mediates parasite protection by blocking complement mediated killing, Factor H activity was depleted in a titratable manner using a polyclonal

70 Factor H antibody. Increased levels of C3b deposition (Figure 2.7A) and increased C5b-9 formation (Figure 2.7B), as well as a corresponding decrease in viability (Figure 2.7C) were observed. Likewise, Factor H depleted serum exhibited elevated levels of C5b-9 on all strains tested (Figure 2.7D). Adding back FH to depleted serum completely suppressed

C5b-9 formation on wild type parasites. Moreover, these data provide further evidence to support the idea that FH directly regulates amplification of the complement response.

Adding back FH did not affect C5b-9 formation on the DKO strain, and only partially suppressed C5b-9 levels on single knockout strains. These results independently confirm that the interactions of SRS29C and SRS25 with FH are critical, as complement activation was not regulated in the absence of these FH binding proteins (Figure 2.7D).

Notably, the effects of blocking or depleting Factor H on C5b-9 levels and on the viability of wild type parasites was only partial and did not reach the significantly elevated levels of C5b-9 and lack of resistance to serum killing observed by the DKO in 10% NHS.

Taken together, these results further establish that additional interactions with C4BP, and possibly other regulators, facilitate WT parasite resistance to serum killing.

2.2.9 Lack of Dsrs25Dsrs29C DKO parasite killing in mouse serum does not contribute to protection in vivo due to low terminal pathway complement activity

In order to determine if the mouse model was suitable for testing complement susceptibility of DKO parasites in vivo, parasites were exposed to fresh mouse serum in vitro and parasite viability was measured by flow cytometry. Compared with the Dsrs25

Dsrs29C DKO parasites exposed to 10% human serum, neither female nor male C57BL6 mouse serum significantly impacted parasite viability (Supplemental Figure 2.9A). This data implied the Dsrs25Dsrs29C DKO parasites would be resistant to serum killing in the

71 murine host. Accordingly, mice infected with the DKO strain did not exhibit differences in survival compared with mice infected with the WT strain (Supplemental Figure 2.9B).

Several studies have confirmed that the low complement activity in laboratory mouse strains is due to low levels of complement proteins, particularly terminal pathway proteins C6, C7, C8, C9 (396, 397). Adding increasing amounts of purified human C3 to mouse serum, up to physiological concentrations found in human serum (1 mg/ml), did not affect parasite viability, indicating that the low levels of murine C3 do not determine the lack of parasiticidal activity in mouse serum (Supplemental Figure 2.9C). Rather, it would appear to be due to the low levels of terminal complex components that likely preclude complement effector activity. Taken together, these results demonstrate that the mouse model is not suitable for testing complement-mediated killing in vivo and requires testing in another animal model replete with complement activity similar to that of human serum. However, despite the lack of parasiticidal activity, several other complement effector functions are still functional in mouse serum that warrant future investigations.

2.3 Discussion

Since the initial study done almost 30 years ago, the parasite factors mediating T. gondii complement resistance have remained elusive. Complement regulation in the host, and for many pathogens, is a complex trait that requires multiple factors. In this study, using forward and reverse genetic approaches, we identified two surface coat proteins

SRS29C and SRS25 that recruit C4BP and FH to downregulate lectin and alternative pathway activation, respectively. Here, we determined that the lectin pathway was a major contributor to C3b deposition, which was in contrast to previous work (173). We also determined a critical role for the AP, consistent with previous work (1). Using single and

72 double knockout parasite strains, we determined the relative capacities of SRS29C and

SRS25 to recruit C4BP and FH, respectively. Based on these interactions, we deduced the contributions of C4BP and FH to parasite resistance to serum killing. Although the lectin pathway largely contributed to parasite recognition and initiation of the complement cascade, it was not a major target for parasite regulation. This is supported by the small impact the loss of C4BP had on the viability of Dsrs29C parasites and C5b-9 formation, possibly the result of C4BP largely targeting C4b, and to a lesser extent, C3b (108).

Because the AP effectively utilizes C3b deposited by CP or LP as a substrate for the AP convertase, a process known as the amplification loop (57, 123), the AP was identified as the most relevant pathway the parasite must inactivate to resist serum killing. This was supported by the loss of serum resistance exhibited by Dsrs25 parasites which failed to block C5b-9 MAC complex formation. Blocking or depleting FH in NHS, but not C4BP

(unpublished observations), further indicated that the AP is the major pathway targeted by the parasite SRS proteins to confer serum resistance. Although SRS25-mediated recruitment of FH was the major factor regulating parasite resistance, it was ultimately the cooperative function of SRS29C and SRS25 to recruit both C4BP and FH that rendered parasites resistant to complement attack. Based on this work, we demonstrate that it is ultimately how these two pathways synergistically work together to mediate their attack that is of fundamental importance. Our approach to use a forward genetic screen facilitated this discovery by identifying two parasite factors that work cooperatively to regulate two arms of an innate sensing pathway in order to confer parasite resistance to serum killing.

By employing such non-assumptive experimental strategies, we identified novel

73 interactions that point to critical points of intervention for therapeutic strategies to render parasites such as Toxoplasma and Plasmodium to be more susceptible to host defenses.

The capacity of T. gondii to recruit C4BP and FH to its surface raises an interesting set of questions pertaining to exactly how the parasite achieves this. Of particular interest is the ability of the parasite to recruit FH, as it played the larger role in resistance to serum killing. Studies have determined the AP utilizes C3b and FH to discriminate between self and non-self surfaces, where FH recognizes ubiquitous host polyanion structures like sialic acid or heparan sulfated proteoglycans (SPGs) (51, 131). Several bacterial pathogens exploit this interaction by either scavenging sialic acid to coat their surfaces or synthesizing their own to recruit FH and thereby protect their surface from complement attack (398–

400) . Several studies have demonstrated that T. gondii possess a lectin like activity associated with its ability to interact with host SPGs (255, 256). Intriguingly, the crystal structure for SRS29B revealed a dimer dependent basic groove capable of docking a host heparan sulfated proteoglycan (265). Because SRS proteins share sequence identity, it is possible that other SRS proteins may exhibit a similar capacity to bind SPGs, as comparative modeling predicted for SRS57 (265). One could speculate that T. gondii utilizes its capacity to bind SPGs as a mechanism for FH recruitment. Based on our study,

SRS29C fulfills two of the criteria for AP regulation: it directly binds C3b and recruits FH, thus it is a strong candidate for structural studies to determine whether it has the capacity to bind SPGs.

Until now, a biological role for the SRS superfamily of proteins has only been inferred and not ascribed. In this study, we identified two SRS proteins that regulate serum resistance by their ability to specifically recruit host regulators to control complement

74 activation. Notably, our forward genetic studies also identified allelic differences that likely play a contributing role in the differential deposition of C3b, which was apparent between

Type I and Type II strains. It is well established that polymorphisms result in expression level differences, as has been observed for a number of the secreted effector proteins that exist as important virulence determinants in the murine model (338, 382). Previous work suggested that high expression of SRS29C was responsible for altering parasite virulence in a mouse model of acute infection, but precisely how SRS29C altered virulence kinetics was not resolved (260, 261). Unlike published Toxoplasma secreted effector proteins that are known to specifically target defined intracellular signaling pathways and cell autonomous immunity that impact parasite pathogenesis, a role for the SRS proteins in regulating either host cells or the innate immune system has only been implied. The ability of two Toxoplasma surface coat SRS proteins to recruit host regulators that inactivate complement C3b to confer serum resistance formally establishes a role for the SRS proteins as virulence factors that promote parasite persistence. It is possible that the limited polymorphism that is present between Type I and Type II alleles regulates the expression level difference that accounts for the strain-specific phenotype, but equally, the polymorphism may impact FH host regulator binding. In support of this, recent studies examining allelic differences in Streptococcus pneumoniae FH binding protein CbpA have shown that these differences contribute to differential levels of FH binding across several strains (401).

In this study, we determined that the absolute levels of C3b between Type I and

Type II strains had no impact on complement-mediated lysis due to the equal capacity of these strains to regulate C3b inactivation. As many pathogens have efficient strategies to

75 resist complement-mediated killing, the complement system has overcome this step by employing several effector functions that are dependent on the generation of C3b and C3a.

This raises interesting questions about the biological relevance of differential C3b deposition between Type I and Type II strains. It is highly likely that the parasite’s ability to regulate the level and the form of C3b may be used to exploit several of these effector functions to modulate virulence. In this study, we determined that the surface of T. gondii is predominantly coated with iC3b, an important effector molecule that is a ligand for several receptors on both myeloid cells and lymphocytes, thus having potentially important implications in pathogenesis. Opsonization with iC3b have been associated with increased phagocytosis for several pathogens (88, 402). Although phagocytosis is not the major mode of T. gondii entry into the cell, recent studies have shown that mouse avirulent Type II strains are preferentially phagocytosed by murine macrophages, whereas mouse virulent

Type I strains are more invasive (308). Complement is also an important proinflammatory mediator, where regulation of complement activation by T. gondii may impact the levels of C3a released in the local environment, impacting recruitment of innate immune cells and cytokine production from these recruited cells. Thus, future work will be necessary to determine specifically how complement regulation affects these critical effector functions and if this impacts T. gondii pathogenesis.

Our studies have also raised interesting questions about the use of the mouse model to study the biological relevance of the SRS proteins in vivo. We generated Dsrs25Dsrs29C

DKO parasites that were highly susceptible to human serum, whereas they remained resistant to lysis in mouse serum. Using higher concentrations of mouse serum resulted in

~60% lysis in up to 80% fresh mouse serum (unpublished observations) but did not

76 contribute to differences in virulence in vivo. Total levels of complement and significant variations in efficiency of complement activity between species are well documented

(403). For example, mouse serum has been shown to have exceptionally low complement activity compared to that of human and rabbit serum (396, 404). Additional differences in sex and among various laboratory strains of mice in their complement activity have also been reported, thus presenting several additional limitations in the study of the protective effects of complement mediated lysis in laboratory mice (397, 405). Thus complement- mediated lysis is not likely the important factor in mediating murine protection in vivo. In addition to low complement activity, differences in species-specific affinities for complement regulatory proteins could also be a contributing factor to these discrepancies.

There is considerable variation in regulatory proteins across mammalian species, with only

60% homology between human and mouse FH. We did not compare recruitment capacities or affinities for murine FH and C4BP using our single and double knockout strains, however it is possible that SRS29C and SRS25 may have variable affinities for mouse complement regulators. It is also possible that SRS29C and SRS25 may not be the relevant

SRS proteins mediating protection in mouse serum. Similar observations have been made with the surface glycolipid LPG in Leishmania which promotes significant resistant to human complement, however plays no role in resistance to mouse complement (406). This lends to the idea that T. gondii has evolved this structurally similar yet semi-redundant family of SRS proteins to fine tune complement regulation in multiple hosts.

Complement evasion is an essential step in the establishment of infection for many microorganisms, which often employ multiple and redundant mechanisms to escape complement killing. For almost 30 years, a critical component of the humoral response to

77 T. gondii has been left almost completely uncharacterized. Here, improved strategies to further characterize the interactions between the complement system and T. gondii lead to significant advancement in understanding the parasite factors that regulate complement activation. In this study, we used forward and reverse genetics strategies to demonstrate that complement regulation was achieved by SRS29C and SRS25, through the recruitment of soluble host regulators FH and C4BP. Using SRS29C and SRS25 single and double knock strains to measure contributions of C4BP and FH to parasite resistance led to a better understanding of roles of the lectin and alternative pathways in response to T. gondii. Our findings support a model in which LP initiation of C3b deposition, which is dependent on

SRS29C expression and the presence of surface carbohydrate ligands, is primarily targeted for inactivation by FH in order preclude an active platform for the AP amplification loop.

However, the cooperative capacity of both C4BP and FH are ultimately critical for parasite survival. Most notably, this work formally demonstrated that strain-specific differences in

SRS genes have an important impact on interactions with the complement system and assigned novel biological function for SRS proteins in regulating innate humoral immunity.

2.4 Materials and Methods

2.4.1 Parasites

Toxoplasma gondii tachyzoites were maintained by in vitro passaging in confluent monolayers of human foreskin fibroblasts (HFF) in Dulbecco modified Eagle medium

(DMEM) with 10% fetal bovine serum (FBS), 2mM glutamine, penicillin, streptomycin and 10µg/ml gentamicin at 37°C, 5% CO2. Parasites were counted using a hemocytometer.

78 2.4.2 Generation of CZ1 Δsrs29C and CZ1 Δsrs25 mutant strains

A clonal isolate of CZ1-Δku80/Δhpt was used to generate the CZ1-Δku80/Δsrs29C and

CZ1-Δku80/Δsrs25 parasites. The SRS29C and SRS25 genes were replaced by the drug selectable marker HPT (hxgprt – hypoxanthine-xanthine-guanine phosphoribosyl transferase) flanked by 40bp homologies to SRS29C and SRS25 using CRISPR/Cas system (SRS29C guide RNA sequence: GCCTACGACACTTATAAGTG and SRS25 guide RNA sequence: GTGTCAAAGTACGTGTGTCC). Fifteen micrograms of HPT repair oligo and 30 µg of guide RNA was transfected into by electroporation using the BTX system. The parasites were then allowed to infect T25 flask of HFFs overnight, after which the medium was changed to mycophenolic acid (MPA) and xanthine at 50ug/ml for HPT positive selection. After two passages, the parasites were cloned using single-cell method into 96 well plates. Δsrs29C and Δsrs25 clones were identified by PCR of the integrated of HPT into the open reading frame using the following primers: (SRS29C forward:

GCCGGAGGGCTTCTCCATCC; SRS25 forward: GACTTGCAGGACCAGTTTC and

HPT reverse: AACCCATTGAAGACTACGGC). SRS29C deletion was further confirmed by staining and analyzed by flow cytometry

2.4.3 Flow cytometry to determine C3b deposition, Factor H binding and parasite viability

Parasites were collected from a freshly lysed monolayer of HFF cells, filtered, and washed twice with PBS to remove media. For time course studies, 1x106 parasites (per time point) were incubated with 10% non-immune human serum (pooled NHS, Cederlane; MBL deficient serum, BioPorto; FH depleted serum, Complement Technologies, Inc) in

++ complement activating buffer HBSS (Hanks Buffered Saline Solution, 1mM MgCl2,

79 0.15mM CaCl2). All serum was tested by IFA and flow cytometry for the presence or absence of human anti-Toxoplasma IgG prior to use. Complement activation was stopped using cold PBS and washed two times with cold PBS to remove excess serum. For viability assays, parasites were exposed to 20%, 40% and 60% NHS, washed three times with cold

PBS to remove excess serum, and stained with a fixable viability stain (Life Technologies) for 15 minutes. Parasites were then fixed with 1% PFA solution for 10 minutes, washed, and stained with mouse α-human C3b/iC3b antibody (1:500) (Cedarlane), goat α -human

MBL (1:1,000) (R&D Systems), mouse α-human C5b-9 (1:500) (Santa Cruz

Biotechnology) or goat α -human Factor H (1:2000) (Complement Technologies), followed by α-mouse APC (eBiosciences) or α-goat APC conjugated antibody (Jackson

Laboratories), both at 1:1,000. For lectin binding assays, 5 ug/ml of biotinylated lectins

(Vector Labs) were incubated with parasites for 1 hour on ice, washed and followed by incubation with SA-APC (1:1000). 20,000 events were collected on a BD Fortessa flow cytometer and analyzed by FACs DIVA, FlowJo and Prism software.

2.4.4 Western blotting

Parasites were treated with 10% NHS for 20 minutes, washed three times with cold PBS, and lysed with 1% NP-40 lysis buffer (1% NP-40, 150mM NaCl, 50mM Tris pH 8.0) for one hour on ice and pelleted at 14,000 rpm for 20 minutes to remove insoluble material.

Lysates were prepared with 2X Laemmli sample buffer (Biorad) with 5% 2- mercaptoethanol and boiled for 5 minutes at 95°C. Samples were loaded onto 12% polyacrylamide gels (Biorad) and transferred onto nitrocellulose using GenScript eBlot system. Membranes were blocked for 1 hour with 5% non-fat dry milk in PBS + 0.05%

Tween-20. The following antibody dilutions were used: rabbit α-SAG1 1:1,000, rabbit α-

80 CRD 1:2,000, rabbit α-C4BP (AssayPro) α-rabbit HRP 1:10,000 (Sigma), goat α-C3b and goat α-Factor H antibodies (CompTech) were both used at 1:20,0000, α- goat HRP 1:5,000

(Santa Cruz Biotechnology, Inc.), goat α-MBL (R&D Systems), mouse α-SRS29C, α- mouse HRP 1:1,000 (Sigma) . Proteins were detected with Clarity Western ECL Substrate

(Biorad).

2.4.5 Factor H blocking and supplementation

10% NHS in HBSS++ was incubated with 1:100 or 1:400 dilution of polyclonal goat anti- human FH for 1 hour on ice, and then added to 1x106 parasites and incubated at 37°C for time course experiments. For FH supplementation, 10% or 40% FH depleted serum was supplemented with 500μg/ml purified FH (Complement Technologies, Inc.) prior to incubation with parasites.

2.4.6 Mouse serum parasite killing assays

WT CZ1 and Δsrs25Δsrs29C DKO parasites were prepared as described above and incubated in either 10% fresh mouse serum or 10% non-immune human serum for 30 minutes at 37°C. For C3 add back experiments, 10% mouse serum (NMS) from a naïve

C3-/- female mouse was supplemented with increasing amounts of purified human C3 and incubated at 37°C for 30 minutes. Parasites were washed twice with cold PBS, stained with a fixable violet viability dye and fixed with 1% PFA at room temperature for 10 minutes.

2.4.7 Statistical analysis

Data were expressed as mean ± standard error of the mean (SEM) of at least three independent experiments. Statistical analyses were performed using Student’s t test between strains or control serum, and mean differences, which are denoted by an asterisk, were considered statistically significant if *p<0.05, **p<0.01, ***p<0.001.

81 2.5 Figures

Figure 2.1. Strain-specific difference in C3b deposition does not affect serum resistance in non-immune human serum. (A) 1x106 Type II ME49 (closed circle) and Type I RH (closed square) parasites were incubated in 10% NHS in HBSS++ over 60 minutes and C3b deposition was measured using flow cytometry as described above; ME49 parasites in 10% heat inactivated serum (Δ56) are used a negative control (open square, dotted line). The data shown are an average of at least three independent experiments done in duplicate and represented as mean ±SEM, ***p<0.001, one-way ANOVA. (B) Western blot showing difference in levels of C3b between 1x106 Type II ME49 versus 1x106 RH parasites incubated 20’ in 10% NHS at 37°C, but no absolute differences inactivated

82 complement iC3b (C) Flow cytometry results from one representative experiment of C3b deposition time course comparing Type II strains ME49 (black) and CZ1 (gray) in 10% NHS over 60 minutes. (D) Western blot analysis of kinetics of C3b inactivation on Type II ME49 parasites over 60 minutes in 10% NHS probed for C3 and its catabolites using a goat α-human C3 polyclonal antibody 1:20,000 (Complement Technologies) and donkey α-goat 1:5,000 (Santa Cruz Biotechnology). Arrows in (D) correspond to sizes of C3 (active C3b a chain 110 kDa, b chain 75 kDa; inactive iC3b a1 68 & a2 43kDa, and C3dg 41kDa). p22 (α-SRS34A, formerly SAG2) 1:2,000 was used as loading control. Western blot images are representative of at least two independent experiments with similar results. (E) Parasite viability and C5b-9 pore formation was determined by exposing parasites to 10%, 20%, 40%, 60% non-immune (NI) or immune (I) serum for 60 minutes at 37°C. 20% heat inactivated non-immune serum was used a negative control. Flow cytometry was used to measure parasite susceptibility by staining parasites with a fixable viability stain (Life Technologies) and (F) membrane attack complex formation using monoclonal mouse α- human C5b-9 1:500 (Santa Cruz Biotechnology). Immune serum was used a positive control for parasite lysis due to classical pathway activation (D, E). Flow cytometry results are representative experiment of at least two independently performed experiments done in duplicate.

83 Figure 2.2. Forward genetics screen and QTL analysis identify several loci regulating C3b deposition. (A) Flow cytometry results for forward genetics strategy measuring differences in C3b deposition of recombinant progeny from a Type I GT1-SNFR x Type II ME49-FUDRR genetic cross. Parental strains and recombinant progeny were incubated in 10% NHS in HBSS++ for 30 minutes. C3b was detected with monoclonal mouse anti- human C3b 1:500 (Cedarlane) and analyzed by flow cytometry. MFI values shown are averages from two independently performed experiments done in triplicate. Data are represented as mean ±SEM. (B) QTL (quantitative trait loci) mapping of the loci on Toxoplasma chromosomes that regulate C3b deposition using J/qtl software with previously generated genetic maps of parental strains and recombinant progeny(382). The LOD scores indicate the log likelihood of association with C3b deposition and are plotted relative to mapped markers across the Toxoplasma genome in numerical chromosome order, with the dashed lines represent the average cutoff for significance across all mapped genes and solid lines represent the parasite locus that maps to a genetic marker with a significant LOD score.

Figure 2.3. SRS29C promotes C3b deposition and is covalently modified by C3b. Flow cytometry results for C3b deposition time course experiments for Type II CZ1 Δsrs29C (A, B) and Type I RH srs29C++ (C,D) compared to CZ1 and RH wild type strains. Figures (A) and (C) are representative of time course experiments of at least three independently performed experiments, which are shown in (B) and (D) as an average of at least 3 experiments done in duplicate for the 30-minute time point. The percentage of binding of C3b was calculated relative to WT strain in (B) and (D). Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, *p<0.05. (E) Western blot analysis of covalent complex formation between SRS29C and heat labile serum protein after incubating 1x107 Type II ME49 parasites with 10% NHS for 30 minutes 37°C. (F) α- C3b immunoprecipitation of 1x107 Type II ME49 parasites incubated in 10% NHS for 30 minutes at 37°C. Parasite lysates used for immunoprecipitation were treated with GPI-PLC for 1 hour at 37°C in order to cleave all GPI-anchored SRS proteins prior to immunoprecipitation. Eluates from the immunoprecipitation were additionally treated with 1M hydroxylamine (NH2OH) to break the covalent complex between C3b and acceptor molecules. GPI-anchored acceptors were probed for with anti-CRD antibodies that recognize the cross-reactive determinant (CRD) epitope that is exposed upon cleavage by GPI-PLC.

Figure 2.4. SRS25 limits C3b deposition. Flow cytometry results for C3b deposition time course experiments for Type II CZ1 Δsrs25 (A,B) and Type I RH Δsrs25 (C) compared to wild type CZ1 and RH strains. Figures (A) and (C) are representative time course experiment of at least three independently performed experiments. (B) is the average of at least 3 experiments done in duplicate at the 30-minute time point. The percentage of binding of C3b was calculated relative to WT strain in (B). Data are represented as mean ± SEM and p-values were obtained using Student’s t-test, ***p<0.001. (D) Western blot analysis of active (C3b) versus inactive forms (iC3b) on Type I RH, RH Δsrs25, and RH SRS29C++ parasite strains incubated in 10% NHS for 20’ at 37°C to show differences in levels of C3dg (arrow, 41kDa). Western blot is representative of at least two independent experiments

Figure 2.5. T. gondii recruits alternative pathway regulator Factor H and lectin pathway regulator C4b-binding protein. 1x107 Type II ME49 parasites were incubated in 10% NHS for 0-60 minutes at 37°C. Western blots of (A) C4BP (rabbit α-C4BP, AssayPro 1:500) and (B) FH (goat α-Factor H, Complement Technologies 1:20,000) binding. Western blots are representative of at least two independent experiments with similar results.

87 Figure 2.6. Serum resistance requires cooperative recruitment of FH and C4BP by SRS25 and SRS29C. (A) Flow cytometry results for parasites incubated in 40% NHS for 10’ and stained with anti-C4BP antibody. (B) Flow cytometry results for parasites incubated in 10% NHS for 30’ and stained with anti-FH antibody. (C) Flow cytometry results for C5b-9 formation on CZ1, CZ1 Δsrs25, CZ1 Δsrs2 Δsrs29C in 40% serum for 30’at 37°C, and as a control DKO parasites in 40% heat inactivated serum (Δ56) NHS. C5b-9 was detected with a monoclonal mouse anti-human C5b-9 mAb (1:500 Santa Cruz Biotechnology). Serum concentrations and incubation times were determined empirically, based on serum concentration that provided the optimal detection of each serum protein, due to differences in abundances in serum. Flow cytometry results for A-C are shown as at least two independently performed experiments. Data are represented as mean ± SEM and p-values were obtained using Student’s t-test, *p<0.05, **p<0.01. (D) Parasites were incubated in 10% NHS for 0-60 minutes and viability was detected using a fixable viability dye (Life Technologies) and measured by flow cytometry. Percent change was calculated relative to parasite viability at 0’. Data are an average of three independently performed experiments.

Figure 2.7. Factor H recruitment enhances parasite resistance to complement- mediated killing. Factor H was blocked by pre-incubating 10% NHS with 1:100 or 1:400 dilution of goat α-human FH pAb for one hour on ice before performing C3b time course experiments. (A) C3b was measured as previously described. Data are shown from one representative time course experiment of two independent experiments (B) C5b-9 formation and (C) parasite viability after 60’ in 10% NHS (black bar), 10% NHS blocked with 1:100 or 1:400 of α-FH antibody (open bars), or 10% heat inactivated serum (gray shaded bar) at 37°C. Data shown are combined averages from two independent experiments done in duplicate. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, **p<0.01. (D) C5b-9 formation was compared on parasites incubated in 40% NHS (black bar), 40% FH depleted NHS (open bar) and 40% FH depleted serum addition of 500 μg/ml purified human FH for 30’. Δsrs29B parasites were used as a control to show specificity. Data shown are SEM of means from two representative experiments done in duplicate.

Supplemental Figure 2.1. Titration of NHS determines 10% is optimal concentration for measuring C3b deposition by flow cytometry. Type II ME49 parasite were incubated in serum serially diluted two-fold, from 40% to 2.5%. Dilutions were made in Hanks ++ Buffered Saline Solution (HBSS ) supplemented with 0.15mM CaCl2 and 1mM MgCl2 (for lectin and alternative pathway activation, respectively). C3b deposition was measured by flow cytometry by staining parasites with a monoclonal mouse α-human C3b/iC3b antibody (Cedarlane) at 1:500. 10mM EDTA inhibits all complement pathways and was used as negative control.

Supplemental Figure 2.2. C3b deposition on parental strains GT1-SNFR (Type I) and ME49-FUDRR (Type II). To determine if GT1-SNFR and ME49-FUDRR were suitable for a forward genetics approach, C3b deposition was measured by flow cytometry. The 2- 3-fold difference between these strains was sufficient to proceed with screening I x II progeny to identify to genetic basis for this difference.

90 Supplemental Figure 2.3. Candidate genes for reverse genetic studies to validate QTLs. (A) List of candidate genes for reverse genetic screens include the following criteria: signal peptides and abundantly expressed tachyzoite SRS surface proteins (toxodb.org). (B) Differential expression of candidate gene SRS29C between Type I (RH, red) and Type II strain (CZ1, gray) measured by flow cytometry. Dotted line represents unstained control (C) Representative non-synonymous (open diamond) and synonymous (shaded diamond) SNPs between ME49 and GT1 in the 5’-UTR and coding regions of the SRS29C locus and (D) SRS25 locus.

Supplemental Figure 2.4. Deletion of SRS29C and SRS25 in CZ1 Type II strain by CRISPR-targeted insertion of HXGPRT gene. (A) Schematic of CRISPR-mediated disruption of SRS29C and SRS25 genes via insertion of the HXGPRT gene by double crossover homologous recombination. Arrows and numbers indicate the primers used to check for HXGPRT insertion with predicted PCR amplicon size. (B) PCR amplification of DNA for wild type CZ1 strain, Δsrs29C, Δsrs25, and Δsrs25Δsrs29C DKO strains. Primers combinations and corresponding sizes are as follows: 1/2 SRS29Cforward/SRS29C reverse (825 bp WT; 2,430 bp Δsrs29C); 1/3 SRS29C forward, HXGPRT reverse (803 bp Δsrs29C ); 4/5 SRS25 forward/ SRS25 reverse (317bp WT; 1,911 bp Δsrs25 ); 4/3 SRS25 forward, HXGPRT reverse (818bp Δsrs25) Bands are consistent with predicted sizes. (C) SRS29C expression for Δsrs29C (orange line) and Δsrs25 Δsrs29C (blue line) strains measured by flow cytometry. Black dotted line represents unstained control.

Supplemental Figure 2.5. SRS25 deletion does not significantly alter expression of abundant tachyzoite proteins SRS29B, SRS34A, SRS29C but not SRS57. CZ1 Δsrs25 parasites were stained with rabbit α-SRS29B (1:5,000), rabbit α-SRS34A (1:5,000), mouse α-SRS29C (1:200) and mouse α-SRS57 (1:500) for 30 minutes on ice followed by goat α- rabbit APC (1:1,000) or rabbit α-mouse APC (1:1,000) for 30 minutes and then analyzed by flow cytometry. Representative histograms of SRS29B, SRS34A, SRS29C and SRS57 are shown for WT parasites (black solid line) and Δsrs25 parasites (red solid line) in the top panel and MFI values are provided in the table in the lower panel.

93 Supplemental Figure 2.6. T. gondii primarily activates the lectin pathway. (A) Flow cytometry results of C3b deposition time course over 60 minutes at 37°C on Type II ME49 tachyzoites incubated in 10% NHS in HBSS++ (closed circle, solid line) or with HBSS++ with 10mM MgEGTA to inactivate LP (open circle, solid line). 10mM EDTA was added to 10% NHS to inactivate all complement pathways (open square, dotted line) Data shown in (A) are a representative time course experiment of at least three independently performed experiments, which are shown in (B) as the average at the 30-minute time point for at least 3 experiments done in duplicate. Data are represented as mean ± SEM and p-values were obtained using Student’s t-test, *p<0.05. (C) Type II ME49 parasites were incubated in 10% serum from MBL deficient serum in HBSS++ (open circle, dotted line), 10% NHS in HBSS++ (solid circle, solid line) or 10% heat inactivated NHS in HBSS++ (open square, solid line). Data shown in C are one representative time course experiment of at least three independently performed experiments, which are shown in (D) as the average at the 30- minute time point. Data are represented as mean ± SEM and p-values were obtained using Student’s t-test, *p<0.05. In (B, D) control sera were arbitrarily set at 100% and the percentage of C3b deposition was calculated relative to control.

Supplemental Figure 2.7. Differential surface carbohydrate composition between Type I and Type II strains affects MBL recognition and is SRS29C-dependent. Carbohydrate ligands on the parasite surface were surveyed using a panel of biotinylated lectins with known specificities (Vector Laboratories, α-mannose/α-glucose (ConA Concanavalin A), N-Acetylglucosamine, GlcNac (WGA, wheat germ agglutinin), galactose (RCA, Ricinus communis agglutinin I) and, N-Acetylgalactosamine, GalNAc (DBA, Dolichos biflorus agglutinin). 1x106 parasites were incubated with 5 μg/ml of biotinylated lectin for 1 hour on ice, followed by streptavidin-APC (1:1,000) for 30 minutes, and analyzed by flow cytometry. Representative histograms of lectin binding between Type II (ME49, blue) (A) and Type I (RH, red) (B) compared to unstained control (black, dotted line) (C) Flow cytometry results of MBL (mannose binding lectin) binding to Type II (ME49 blue) and Type I (RH red) parasites incubated in 10% NHS for 60’ on ice. MBL was detected using goat α- human MBL (R&D Systems 1:1000). Data shown are an average of at least two experiments done in duplicate. Error bars represent SEM. (D) Representative histograms of lectin binding for CZ1 (gray, open) and CZ1Δsrs29C (orange, shaded) parasites and (E) CZ1Δsrs29B (shaded, brown). Unstained control is shown in black dotted line. (F) Flow cytometry results for MBL binding of CZ1 (gray) and CZ1Δsrs29C (orange) parasites incubated in 10% NHS for 60’ on ice. Data shown are an average of at least three experiments done in duplicate. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, ***p<0.001.

95 Supplemental Figure 2.8. SRS25 SRS29C double deletion does not significantly alter SRS29B, SRS34A or SRS57 expression, but does impact C3b deposition. (A) Δsrs25Δsrs29C DKO parasites were stained with α-SRS29B, α-SRS29C, α-SRS34A, and α-SRS57 as described in Supplemental Figure 4. Representative histograms of SRS29B, SRS34A, SRS29C and SRS57 staining are shown for WT parasites (black solid line) and Δsrs25Δsrs29C DKO parasites (blue solid line) in the top panel and MFI values are provided in the table in the lower panel. (B) DKO parasites (blue dotted line) and WT parasites (gray solid line) were incubated in 10% NHS over 60 minutes at 37°C and C3b deposition was measured by flow cytometry. DKO parasites exhibited a significant increase in C3b deposition. (C) An average of at least 3 experiments done in duplicate for C3b deposition of WT (gray bar) and DKO parasites (blue bar) at the 30-minute time point. The percentage of binding of C3b was calculated relative to WT strain in. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, **p<0.01.

96 Supplemental Figure 2.9. Low complement activity in mouse complement precludes killing of Δsrs25 Δsrs29C DKO parasites in vivo. (A) Complement activity of mouse serum versus human serum. WT CZ1 and Δsrs25Δsrs29C DKO parasites were incubated in either 10% fresh mouse serum or 10% non-immune human serum for 30 minutes at 37°C. Parasite viability was measured by flow cytometry using a fixable violet viability dye. Results shown are an average of two independent experiments. (B) 2x103 Type II CZ1 parasites (black solid line) or CZ1 Δsrs25Δsrs29C parasites (blue dotted line) were injected i.p. into 5 female C57BL6 mice and survival was monitored. (C) 10% mouse serum (NMS) from a naïve C3-/- female mouse was supplemented with increasing amounts of purified human C3 to demonstrate that low complement activity in mouse serum was not at level of C3.

97 CHAPTER 3: TOXOPLASMA SURFACE COAT PROTEIN SRS29C

RECRUITMENT OF FACTOR H TO EVADE COMPLEMENT AND

REGULATE PARASITE VIRULENCE IS SULFATED PROTEOGLYCAN (SPG)

DEPENDENT2

3.1 Introduction

Toxoplasma gondii is a widespread intracellular protozoan pathogen that can infect a wide range of hosts, including humans, and establishes a life-long chronic infection. The initial steps of infection are dynamic processes that require multiple interactions with host factors and modulations of innate immunity that together ultimately facilitate the establishment of a successful infection. Pathogen surface proteins are at the forefront of these interactions. T. gondii’s surface coat is dominated by a family of structurally related proteins known as the SRS (SAG1-related sequence) superfamily (383) that promote attachment to host cell surfaces prior to their invasion. Toxoplasma uses this developmentally-expressed yet antigenically distinct repertoire of surface adhesins to initiate a concerted chain of events to gain entry into any nucleated cell (265, 383). Several

SRS proteins, including SRS29B and SRS57 (formerly SAG1 and SAG3, respectively) have been associated with attachment to host cells, and this trait has been attributed to their capacity to bind ubiquitously expressed host heparan sulfated proteoglycans (SPGs) (255,

258, 385, 407–409). Proteoglycans consist of a core protein with one or more attached

______2Citation: Sikorski, P.M., Pszenny, V., Sardinha da Silva, A., Khan, A., Gray, T., Commodaro, A.G., Kennard, A., Boulanger M.J., and Grigg M.E. Toxoplasma surface coat protein SRS29C recruitment of Factor H to evade complement and regulate parasite virulence is sulfated proteoglycan dependent. In preparation.

98 linear chains of highly anionic polysaccharides, known as glycosaminoglycan (GAG) chains (410). Heparan sulfates are a type of GAG chain that are abundantly and ubiquitously found on cell surfaces and extracellular matrices (411). Structural studies of the major surface antigen SRS29B have identified a conserved dimeric structure exhibiting a positively charged groove that can accommodate anionic SPGs (265, 412). SPGs have been shown to contribute to bacterial and viral pathogenesis by facilitating host cell entry and dissemination (413), however experimental evidence for the role of SPGs during acute

T. gondii infection in vivo is lacking.

The SRS superfamily is also thought to mediate several additional biological functions crucial to establishing infection, including modulation of innate immunity (261,

269, 414), and activation of the adaptive immune response (268). Expression of SRS proteins during acute infection has a dual role in acute infection as they are also thought to regulate virulence in order to promote parasite persistence (414, 415). SRS29B and

SRS34A are highly immunogenic tachyzoite surface antigen that elicit high antibody titers during acute infection, which is assumed to regulate virulence by targeting the immune response against the tachyzoite stage to control parasite proliferation (268, 364, 368, 383).

Immunomodulatory roles for SRS proteins during acute infection have also been described.

SRS29B has been associated with inducing a strong inflammatory response in vivo (269), whereas recent studies have shown that SRS29C, a differentially expressed protein between mouse-virulent Type I and -avirulent Type II strains, attenuates acute virulence

(261). The immune factors or host ligands in which these SRS proteins interact with to elicit such differential responses, however, remain poorly understood.

99 While facilitating initial interactions with the host, T. gondii in the extracellular environment is vulnerable to attack by humoral factors, such as the complement system.

The complement system is a critical component of humoral immunity that all invading pathogens must overcome to establish infection. This first line of defense effector system activates a potent cascade of proteolytic events on the surface of invading pathogens, leading to opsonization with the central protein C3b and direct lysis. While classical and lectin pathways are responsible for detecting pathogens and initiating complement activation, the alternative pathway (AP) is always on at low levels and has the capacity to potentiate a rapid amplification loop. Thus the alternative pathway is stringently regulated by host regulator protein Factor H (FH) to breakdown any deposited C3b and prevent cell damage and inflammatory responses (391). AP regulatory protein FH recognizes host sialic acids and heparan sulfated proteoglycans to discriminate between host and foreign surfaces

(50, 120, 121). Thus, many pathogens often encode either molecules that mimic or sequester host “self” molecules to recruit complement regulators as a means to down regulate complement activation (416–419) and overcome complement attack.

Studies in Chapter 2 established that SRS29C is not only an acceptor molecule for

C3 deposition, but it is also a regulator of complement system activation through its interaction with host regulator proteins Factor H (FH) and C4b-binding protein (C4BP).

Regulation of the alternative pathway by FH, however, was more critical in protecting parasites from serum killing. Both C3b and FH were bound by SRS29C. It is known that host SPGs increase the affinity of FH for C3b in order to discriminate self from non-self

(121). Given the capacity of T. gondii to interact with SPGs and the role of SPGs in the recruitment of complement host regulators, we hypothesized that T. gondii’s interaction

100 with host SPGs facilitates the recruitment of FH. Here, we used a structural approach to gain further insight into the molecular mechanisms underlying complement regulation and the immunomodulatory roles of SRS29C in an animal model of acute toxoplasmosis. We engineered strains transgenically over expressing WT SRS29C and SRS29C with mutations in its highly basic apical surface binding groove in order to investigate whether

SRS29C recruits Factor H in a SPG-dependent manner. Moreover, our in vivo findings demonstrate that a complex interaction between SRS29C, C3, FH and SPGs determines complement regulation and the pathogenesis of Toxoplasma infection in mice.

3.2 Results

3.2.1 SRS29C crystal structure reveals a binding groove lined with basic residues at the apical surface

Previous work showed that SRS29C has a dual role as a C3 acceptor molecule and recruiter of host regulators Factor H and C4BP to the parasite surface to inactivate C3b.

Further, that FH was the critical factor mediating protection from serum killing (Chapter

2). However, the precise molecular mechanisms mediating FH recruitment to the parasite surface were enigmatic. Recognition of host cells by FH is facilitated by the presence of abundantly expressed “self” molecules that associate with host cell surfaces, such as sialic acid and heparan sulfated proteoglycans (HSPGs) (51, 130, 131) . The capacity for some

SRS proteins to bind SPGs indicates a possible mechanism for FH recruitment. Factor H is a single chain glycoprotein consisting of 20 homologous domains known as complement control proteins (CCPs). Factor H typically identifies and binds host cell surfaces through the recognition of C3b and sialic acid or SPGs (51). Factor H has several C3b and SPG binding sites. Domains 1-4 bind C3b and mediate its cofactor activity, with an additional

101 binding C3d site at domains 19-20 that overlaps with the heparin binding domain (121,

124, 127, 128, 130, 420, 421). The structural basis for FH binding has been shown to require the dual recognition of a negatively charged structure and the C3d by FH domains

19 and 20, respectively (130, 421). Our previous worked showed that SRS29C was bound by C3b and strains deficient in SRS29C failed to recruit FH (Chapter 2). We hypothesized that SRS29C specifically interacts with FH to facilitate the catalytic degradation of C3b to iC3b bound to SRS29C. Thus, SRS29C was prioritized for structural analysis.

Sequence analysis revealed that SRS29C harbors a signal peptide followed by two family eight SRS domains (D1 and D2) (261) each containing six cysteines predicted to form three disulfide bonds, and a GPI anchor site (Figure 3.1A). To ensure correct disulfide bond formation in the recombinant protein, an insect cell-based expression system was used. Both SRS29C alleles from the Type I (SRS29C-I) and II (SRS29C-II)

Toxoplasma lineages, which differ in only four amino acid residues, were recombinantly produced and purified as monomers, as determined by size exclusion chromatography and dynamic light scattering. To establish the global architecture of the SRS29C variants, we then determined the crystal structures to 1.84 Å (SRS29C-I) and 2.35 Å (SRS29C-II) resolution. The amino acid differences map to loop structures on D1 and D2 and a central

D2 beta-strand, but do not affect the overall structure or inter-domain organization (Figure

3.1B) consistent with the observation that SRS29C-I and -II have the same effect on attenuating T. gondii virulence (261). Since the SRS29C-I structure was of higher resolution, it was used for further analyses, unless otherwise stated.

SRS29C crystallized as a monomer and was highly ordered enabling the modeling of all residues from Glu61 of D1 through Gly311 of D2 and seven ordered expression tag

102 residues at the C-terminus. Structural analysis revealed tandem canonical SRS domains formed by a beta-sandwich stabilized by three disulfides and linked together by a short linker (Figure 3.1B) (265, 422, 423). Notably, SRS29C is the only characterized SRS protein with a linker that incorporates a defined secondary structure component (Figure

3.1B). This feature has the potential to limit the relative conformational flexibility between the SRS domains, which may facilitate SRS29C in forming multimers on the parasite cell surface. The most intriguing feature, however, is the presence of an extended, contiguous patch of basic residues, comprised of lysines 10, 16, 17, 89 and arginine 14, located on the side of the apical surface of D1 (Figure 3.1C). Structural alignments reveal that this cluster of basic residues maps to an apical groove formed at the homodimeric interface of the related SRS16C structure. Notably, the basic groove of SRS16C is predicted to coordinate negatively charged ligands such as SPGs and modeling studies have shown that the groove can accommodate a heparin chain. It is noteworthy, however, that no definitive binding partner for an SRS protein has been experimentally validated.

3.2.2 An engineered SRS29C dimer specifically binds heparin

The propensity for SRS proteins to dimerize (265, 422, 424) which appears to be necessary for the formation of the basic apical groove, is likely greatly facilitated by the reduced entropic penalties associated with GPI anchor attachment to the parasite membrane. Thus, to support functional characterization of an SRS29C dimer we engineered the insect cell expression vector to encode a C-terminal leucine zipper homodimerization domain (Figure 3.2A). The SRS proteins appear ideally suited to this type of C-terminal modification as the leucine zipper helix should spatially approximate the GPI anchor and therefore not perturb overall structure. Size exclusion chromatography

103 confirmed that the recombinant homodimeric SRS proteins migrated as expected (Figure

3.2B). Since the crystal structure of SRS29B and SRS16C have been previously reported, these two SRS proteins were also produced as homodimers to be used as controls in functional assays.

The localization of the SRS proteins on the exterior of the parasite surface combined with the highly basic apical surface of SRS29C pointed to negatively charged carbohydrates as potential binding partners. To test this, heparin-agarose beads were used to pull down monomer and dimer forms of SRS29C, SRS29B and SRS16C. Only SRS29C was pulled down with the heparin beads and only in the engineered dimer form (Figure

3.2C). A dose-dependency of the SRS29C dimer-heparin interaction was confirmed by native gel electrophoresis, with increasing molar ratios of heparin resulting in a complex with reduced migration consistent with the binding of multiple SRS29C dimers to the same heparin strand (Figure 3.2D). No significant shift in migration was observed for SRS29C monomer even at the highest molar ratio of heparin (Figure 3.2D). To establish that the observed interaction was charge dependent we utilized heparin lacking N-linked sulfates, which showed no binding to the SRS29C by native gel shift (Figure 3.2E). Further, we also tested binding to heparan sulfate, which is a naturally-occurring sulfated glycosaminoglycan with a similar structure to heparin but with reduced charge density.

Highlighting the charged nature of the interaction, a minor but reproducible shift of the

SRS29C dimer was observed in the presence of heparan sulfate (Figure 3.2E).

Interestingly, it has been previously proposed that high levels of sulfation on heparan sulfate, which would mimic heparin, might play a role in implicating particular cell types for infection by apicomplexan parasites (425). Collectively, these data support an

104 interaction between SRS29C and heparin that requires dimerization, is selective relative to related SRS proteins and is charge dependent.

With the high-resolution crystal structure of SRS29C, we were able to generate a panel of strategic mutants designed to probe the most likely heparin binding regions. A total of three lysine residues were targeted for mutagenesis (K10, K16 and K17) because they line the predicted groove identified (Figure 3.2F). Each mutant was recombinantly produced in insect cells and showed the same behavior in solution as native SRS29C. Using native gel shift assays as the readout, mutation of Lys10 to a glutamate, completely abolished binding consistent with the requirement for a charge dependent interaction

(Figure 3.2G). At the periphery of the groove, a double mutation of lysines 16 and 17 to serines also led to complete abrogation of heparin binding (Figure 3.2G). Furthermore, this data demonstrated the capacity of SRS29C to bind SPGs, thus providing direct evidence for an SRS29C-SPG interaction that likely exists as a critical component required for FH recruitment.

3.2.3 SRS29C recruits Factor H through interactions with SPG

To test the hypothesis that FH recruitment was facilitated by the capacity of

SRS29C to bind SPGs, RH transgenic strains that overexpressed either wild type SRS29C

(RH SRS29C++) or SRS29C with the K16/17S mutations (RH SRS29C++ K16/17S) were engineered. The parental strain (RH∆) used for this experiment possesses a disrupted hypoxanthine-xanthine-guanine phosphoribosyl transferase (HXGPRT) gene. We hereafter refer to the RH∆ strain as wild type (WT). The SRS29C coding region was isolated from genomic DNA of the Type II strain 76K or amplified from the pAcGP67B vector with cloned SRS29C that underwent site-directed mutagenesis and cloned into an expression

105 plasmid that contains the SRS29B promoter and un-translated region, the SRS29B signal peptide and GPI anchor. The levels achieved by the SRS29B promoter allowed SRS29C to be expressed to an equivalent degree of the endogenous levels of SRS29C in the majority of the non-virulent strains.

The resulting plasmid was then introduced to WT parasites. The parasites were also co-transfected with a plasmid that allows expression of GFP and Luciferase for bioluminescent and fluorescent imaging experiments. Only clones that possessed SRS29C expression levels comparable to that of the non-virulent Type II strains were selected. The frequency of random integration at various chromosomal sites in Toxoplasma is sufficiently high that insertional mutagenesis is a genetic method of choice in this organism. The probability that the SRS29C expression constructs integrated at the same site is therefore highly unlikely for clones that are derived from independent transfections.

Consistent with this, variation in the expression levels among the independently derived clones was observed. Polar effects due to the site of integration for the expression constructs can likely be excluded as an alternative interpretation of our results given that multiple independent clones produced the same relative phenotype

Next, FH binding was measured by flow cytometry to test if the loss of SPG binding affected recruitment. SRS29C++ parasites incubated in 10% NHS exhibited significantly increased levels of FH on the parasite surface compared to the WT RH strain, however

SRS29C++ K16/17S lost the capacity to recruit FH (Figure 3.3A, 3.3C). Interestingly, these mutations acted in a dominant negative fashion, where the level of FH was below that of the wild type RH, which expresses SRS29C weakly [2, 30]. FH recruitment is also dependent on the presence of C3b [19]. To rule out the possibility that the mutations

106 affected the levels of C3b deposited onto the parasite’s surface, C3b deposition was also measured by flow cytometry. SRS29C++ and SRS29C++ K16/17 strains equivalently promoted increased C3b deposition compared to wild type (Figure 3.3B, 3.3D). However, flow cytometric assays cannot discriminate between active and inactive C3b. Thus,

Western blot analysis for C3b inactivation was performed and showed that the SRS29C++

K16/17 strain exhibited less inactivation product iC3b than SRS29C++, which is consistent with less FH recruitment (Figure 3.3E). These data demonstrated that the lysine residues in the basic groove of the SRS29C homodimer did not affect levels of C3b complement deposition but did result in less Factor H recruitment and thus impacted the degree of C3b inactivation.

3.2.4 Trans-over expression of SRS29C attenuates virulence

Transgenic over expression of either the Type I or Type II allele of SRS29C in a lethal Type I background was previously shown to attenuate acute virulence in the murine model (261). During acute infection, tachyzoites induce a strong Th1 immune response, which is characterized by increased production of IL-12 by innate cells which stimulate

IFN-γ production by T cells (315, 358). This response is influenced by parasite genotype, where infections with Type I strains are thought to be lethal due to dysregulated Th1 cytokines, leading to immunopathology and death (342, 426). From these previous findings, however, one cannot distinguish if the SRS29C over expression effect on virulence is a function of parasite burden as a result of defective parasite replication, or the induction of a more controlled immune response.

To first address if there were any defects in in attachment and invasion in vitro, we utilized luciferase (LUC) expression to quantify successful attachment/invasion events

107 over a time course invasion assay. LUC expressing clones and the parent cell line were inoculated into 24-well plates containing human foreskin fibroblast (HFF) cells. The parasites were allowed to invade HFF cells for different time periods after which extracellular parasites were washed off. Luciferin was added and the plates were imaged using an IVIS-100 bioluminescence imager. Figures 3.4A and 3.4B show that there was an equivalent number of photons detected between WT and transgenic parasites suggesting that there is no difference in attachment/invasion between the parent cell line and SRS29C overexpression clones at time points 15 and 45 minutes. Later time points post-invasion measured differences in growth rate. There was no difference between SRS29C over expresser and the parent cell line at 15- and 24-hours post-inoculation (Figures 3.4A,

3.4B). Toxoplasma is also known invade myeloid lineage cells in vivo so we performed the same assay with the macrophage-like J774 cell line and found that the results were comparable to those obtained using HFF cells (Figures 3.4A, 3.4B). Overall, the results suggest that overexpression of SRS29C has no detectable effect on the ability of parasites to grow, attach and/or invade cell lines in vitro.

To determine whether the previous work (261) demonstrating the attenuation of virulence in vivo was due to an intrinsic difference in the immune response, we employed a co-infection model. To test whether trans-overexpression of SRS29C protects mice from a lethal dose of wild type RH parasites, two groups of CD-1 mice were infected. The first group was infected with 50 tachyzoites of only WT RH GFP-LUC parasites, and the second group was infected with a mix of 50 tachyzoites of wild type RH GFP-LUC positive parasites and 50 parasites SRS29C over expressing “dark” parasites that were GFP-LUC negative. Parasite burden of the wild-type RH parasites was monitored via bioluminescent

108 imaging. Remarkably, by day 7 post infection, wild type parasite burden in the co-infected animals was significantly lower than those infected with RH alone (Figure 3.4C). These results indicated that SRS29C over expression could control wild type RH parasite burden in trans which failed to expand to a level contributing the host death. Mice were not scarified at day 7 to evaluate the levels of the “dark” SRS29C++ parasites in these mice, however previous experiments have shown that mice infected with the SRS29C++ strains regulated parasite burdens and contributed to survival (261). Further experiments would be required to determine if this was an immunologically driven event.

3.2.5 SRS29C-mediated regulation of acute virulence in the lethal Type I background is SPG-dependent

To investigate if loss of SPG binding and FH recruitment in the SRS29C++ K16/17S strain impacted parasite expansion, induction of host immunity and acute virulence, groups of CD-1 mice were injected in the footpad with 25 WT, SRS29C ++, or SRS29C ++ K16/17S tachyzoites and survival was monitored (Fig. 3.5A). As expected, mice infected with wild type parasites quickly succumbed to infection whereas SRS29C++ parasites significantly prolonged survival. Bioluminescent imaging to monitor in vivo parasite burden showed wild type infected mice had significantly higher parasite burdens than SRS29C++ infected mice 9 days post infection, which is consistent with previously published reports (261).

Interestingly, infection with SRS29C++ K16/17S completely abrogated the protective effect of SRS29C overexpression, restoring both virulence and parasite burdens to WT levels

(Figures 3.5A, 3.5B). These data indicate that SRS29C-SPG interactions are critical for host protection.

109 3.2.6 WT SRS29C expressed in Type I strains alters early cytokine responses

To determine if SRS29C over expression impacted virulence by altering the early immune response, the induction of Th1 cytokines present in infection serum were evaluated in mice infected with 25 tachyzoites of RH and the SRS29C++ transgenic strains intraperitoneally. Day 4 was empirically chosen to study this effect, because there is no difference in the expansion of all strains in vivo at this time point. For all three stains, parasite burdens in the peritoneal cavity and spleen were not significantly different, indicating that differences in the cytokine response were unlikely to be directly attributable to differences in parasite load or dissemination (Figures 3.6A, 3.6B). Serum levels of IFN-

γ, IL-12p40, and IL-18 were significantly reduced in mice infected with SRS29C++ compared with the WT strain, whereas those infected with SRS29C++ K16/17S parasites exhibited cytokine levels that were comparable to those produced in WT infected mice

(Figures 3.6C, 3.6D). Taken together, these data suggest that an altered immune response induced by SRS29C overexpression is dependent on the ability of SRS29C to interact with host SPGs. Furthermore, the restoration of the cytokine response in the mice infected with

SRS29C++ K16/17S is not readily attributable to a failure to regulate the early cytokine response in the absence of specific interactions with particular host ligands. Whether this phenotype is dependent on the capacity of SPGs to interact with FH, however, could not be assessed using this experimental set-up.

3.2.7 SRS29C requires complement protein C3 to regulate virulence in vivo

The attenuation of virulence and alteration in lethal cytokine responses as a result of transgenic over expression of SRS29C in a virulent Type I background suggested that high expression of SRS29C in avirulent Type II and Type II strains is an important factor

110 for regulation of parasite virulence. To test this hypothesis, we took advantage of a previously generated Δsrs29C strain in a Type II, mouse avirulent, background (Chapter

2) to determine if loss of SRS29C increased virulence and the inflammatory cytokine production. C57BL/6 mice were infected intraperitoneally with 2x103 wild type, a dose that allows all mice to survive acute disease (death within first 20 days) against the Δsrs29C strains and parasite load, cytokine responses and survival were monitored. In the absence of SRS29C expression, the Δsrs29C parasites were more virulent, killing mice more rapidly, whereas WT infected mice survived significantly longer (Figure 3.7A).

To test whether the complement binding properties of SRS29C were primarily responsible for the altered parasite pathogenesis, C3-/- mice were infected with 2x103 wild type and Δsrs29C parasites. C3-/- animals demonstrated similar mortality kinetics to wild type mice infected with Δsrs29C parasites, where C3-/- mice were more susceptible to infection than C3 sufficient mice infected with the wild type parasite strain. In the absence of C3, wild type and Δsrs29C infected animals exhibited no difference in survival, indicating the parasites induced a similar response when both SRS29C and C3 were absent

(Figure 2.7B). Taken together, these findings show that C3 deficient mice phenocopied the infection kinetics and pathology induced in WT mice infected with the Srs29C- deficient Type II strain, suggesting that SRS29C-mediated protection is C3-dependent.

To rule out the possibility of this phenotype was an artefact of an anomalous clone, we tested another Δsrs29C generated in the Type II Prugniaud (Pru) strain. Mice infected with Srs29C- deficient parasites were likewise more virulent (Supplemental Figure 3.1A).

Here, we also observed that mice infected with Pru Δsrs29C parasites exhibited higher brain cyst burden (Supplemental Figure 2.1B). Histological studies showed increased

111 inflammatory infiltrate and the presence of tachyzoites in the brains of mice infected with

Δsrs29C parasites (Supplemental Figure 3.2C). Together, these data demonstrate that the increase in virulence observed in mice infected with Δsrs29C is associated with increased inflammation and pathology, however further studies are necessary to determine the cellular basis contributing to this phenotype.

3.2.8 SRS29C alters early cytokine responses in vivo

To further investigate the factors contributing to the increased virulence in mice infected with Δsrs29C parasites, both parasite burden and cytokine production were evaluated 6 days post infection. Day 6 was empirically chosen to study this effect, because there is no difference in the expansion of these strains in vivo at this time point. No differences in parasite burdens between WT and Δsrs29C infected animals were observed in the peritoneal fluid, spleen or lungs by counting plaque-forming units (PFUs) (Figure

3.8A), suggesting differences in virulence were not the result of an unregulated proliferation of one parasite clone over the other.

To evaluate the early immune response, systemic cytokine production from serum and local cytokine production from ex vivo splenocyte cultures was measured at day 6 post infection. Δsrs29C infected mice exhibited significantly elevated levels of IL-12 and IFN-

γ in both serum and ex vivo splenocyte cultures (Figures 3.8B-E). Furthermore, ex vivo splenocyte cultures exhibited significantly higher levels of TNF-α (Figure 3.8F), which was not detectable in serum (data not shown). Taken together, these data imply that loss of

SRS29C expression results in a significant increase pro-inflammatory cytokine production that is not unlike the dysregulated Th1 response responsible for acute mortality in mice infected with lethal Type I strains.

112 3.2.9 C3 regulates local cytokine responses in vivo and in vitro

Given the role of C3 in SRS29C-mediated protection, parasite burdens and cytokine responses in C3-/- infected animals were evaluated. C57BL/6 and C3-/- were infected intraperitoneally with 2x103 wild type CZ1 tachyzoites and parasite burden in the peritoneal fluid and spleens was determined by counting the number of PFUs per organ 6 days post-infection. No significant difference in parasite burden was observed between wild type and C3 deficient animals in the peritoneal fluid (Figure 3.9A) and spleens of

C3-/- mice (Figure 3.9B). To determine if a similar dysregulated cytokine response was observed, systemic and local cytokine responses were assessed. No significant differences in IL-12 cytokine levels were observed in serum and ex vivo splenocyte cultures (Figures

3.9C, 3.9G). However, a discrepancy in cytokine production between serum and ex vivo splenocyte cultures for several cytokines was observed. In wild type animals, higher levels of IFN-γ, IL-6, and IL-18 were observed in the serum (Figures 2.9D-2.9E), whereas in ex vivo splenocyte cultures, wild type infected animals exhibited reduced IFN-γ and IL-6 production (Figures 3.9H, 3.9I). The significant increase in cytokine production by ex vivo splenocytes of C3-/- mice is reflective of the phenotype observed in wild type mice infected with Δsrs29C parasites. However, it is not immediately clear why serum cytokine levels were not consistent with those produced by splenocytes. Several studies have indicated that the role of complement in serum is functionally different from that in a local microenvironment of the spleen, whereby Th1 responses are regulated by local C3a and

C5a production by immune cells (152). Several studies have shown that in C3-/- mice, C5 can be activated in the absence of C3 by thrombin to produce C5a and C5b (427). To test this if this was occurring in our system, C3a and C5a levels in the serum of wild type and

113 C3 deficient mice infected with Type II strain ME49 were measured. We observed that C3-

/- mice, as expected, had no C3a present in serum (Supplemental Figure 3.2A), however levels of C5a were similar to wild type mice, indicating a C3 bypass had occurred

(Supplemental Figure 3.2B). These results suggest that C5a is a likely effector molecule that could be mediating the discrepancies in local cytokine production. However, further evaluation of the levels of C5a being produced locally by splenocytes is required to determine if the production in C5a by C3-/- mice is greater than splenocytes from wild type mice, leading to increased cytokine production.

From these findings, it is not clear which complement effector function is important in controlling host protection, as C3a and C3b have distinct effector functions. In Chapter

2, we demonstrated that WT and Δsrs29C parasites are equally resistant to complement mediated lysis despite lower levels of FH in the absence of SRS29C, thus it is not likely that complement-mediated lysis contributes to host protection. Therefore, the C3- dependent phenotypes must be attributed to one or more of the complement system’s diverse effector functions. C3b is an important effector protein that mediates several important functions, including complement-mediated phagocytosis by engaging complement receptors. Several studies have observed dampened proinflammatory cytokine production in response to iC3b coated cells (428, 429). Studies done in Chapter 2 confirmed previous work that has shown T. gondii efficiently inactivates C3b into iC3b

(173)

In order to determine if parasites coated with iC3b influenced cytokine production, parasites in the presence and absence of 40% non-immune human serum (NHS) were incubated with murine bone marrow derived macrophages in vitro. Cytokine and

114 chemokine production 20 hours post infection was assessed by a Luminex assay. Parasites in the absence of human serum produced higher levels of all measurable cytokines and chemokines assayed, including IL-6, MCP-1, MIP-1α, MIP-1β, MIP2, and KC

(Supplemental Figure 3.3A), despite exhibiting similar infectivity (Supplemental Figure

3.3B). One aspect of this interaction with BMDMs that could be altered in the presence of serum is host cell entry. One limitation of using flow cytometry assays to measure infection is that they cannot distinguish between invasion and phagocytosis. Thus, the contributions of differential entry to this response could not be evaluated using this experimental approach. Although it is unclear from this data how iC3b mediates these effects, these data suggest T. gondii may exploit its ability to regulate complement activation to alter local inflammatory responses. Additional studies need to be done to determine the relative contributions of iC3b and anaphylatoxins to host protection.

3.3 Discussion

Toxoplasma gondii employs large families of proteins with diverse effector functions to cooperatively modulate the Th1 immune response in favor of parasite persistence. Our data suggest a complex interaction with several host molecules likely drives the immunomodulatory nature of SRS29C-mediated attenuation of virulence in the mouse model. Structural studies of SRS29B and comparative modeling have supported ligand binding roles for SRS proteins (265, 422, 423). Because of the abundance and ubiquity of SPGs across many different host cells and species, it is believed the main role during T. gondii infection is to facilitate tachyzoite binding to many different host cell types in a large number of mammals (407) to initiate the invasion process. Several studies have determined that a loss of SPGs on several cell types resulted in a significant reduction in

115 T. gondii attachment and infection in vitro (255, 407). In this study, the structural analysis prediction that the basic SRS29C groove supported binding of SPGs was confirmed using sulfated proteoglycan pull down assays and a native gel shift assay.

Heparan sulfated proteoglycans are exploited by many different pathogens to facilitate initial attachment and to subvert host immunity, establishing that this is an important factor in microbial pathogenesis (430). The role of SPGs in T. gondii attachment to host cells in vitro is not exclusive to SRS proteins (264, 431), suggesting functional redundancy and synergy across several families of T. gondii proteins. In this work, we established that SRS29C binds SPGs, however we did not observe any differences in in vitro growth or attachment of Δsrs29C (unpublished observations) or RH SRS29C++ to host cells. Thus, the role of SPGs in mediating attachment was not likely a factor contributing to differences in virulence exhibited by mice infected with these parasite strains. In vivo studies showed no differences in parasite burden between RH, RH SRS29C++ and RH

SRS29C++ K16/17S strains 4 days post infection, suggesting SRS29C-SPG interactions do not play a role in parasite dissemination or infectivity during the initial stage of infection.

Similarly, no differences in parasites burden were observed for wild type infected and

Dsrs29C infected mice at day 6 post infection. Most importantly, the in vivo phenotypes observed for SRS29C supported an immunomodulatory role. Previously, we observed an attenuation of virulence in the case of SRS29C overexpression in a lethal Type I strain

(261), and in this study a corresponding increase in virulence was observed with SRS29C deletion in Type II strains. Notably, co-infection with wild type RH parasites and SRS29C transgenic over expressing parasites reduced parasite burden in trans, further supporting an intrinsic capacity of SRS29C to alter the immune response. Consistent with this role,

116 loss of SRS29C expression resulted in increased inflammation in the brain of surviving mice. Lastly, studies done in Chapter 2 expanded the role of SRS proteins to include regulation of the complement system. SRS29C was implicated in complement regulation by recruiting FH and C4BP to down regulate complement activation, further solidifying the immunomodulatory capacity of this protein.

Host survival relies on a critical balance between inducing potent immune responses in order to control pathogen proliferation as well as the resolution of inflammation to prevent lethal immunopathology (432). Aberrant Th1 responses during acute T. gondii infection with Type I strains leads to immunopathology and mortality, whereas Type II strains are better able to fine-tune this powerful immune response in mice to support host fitness and parasite persistence (342). In this chapter, we determined that mutations which abolish SRS29C binding of SPGs and loss of SRS29C expression abrogated the protective effect of SRS29C expression in vivo. We observed that mice infected with Dsrs29C and SRS29C++ K16/17S parasites exhibited higher levels of Th1 cytokines IL-12 and IFN-g production. It is tempting to speculate that T. gondii strains that highly express SRS29C (i.e. Type II and Type III) exploit their interactions with host SPGs to regulate virulence by modulating or resolving pro-inflammatory processes. In support of this idea, mice infected with SRS29C++ K16/17S parasites were unable to resolve or regulate inflammatory responses compared to animals infected with RH SRS29C++. It is possible that changes in SPG-binding can impact host cell signaling pathways. For example, syndecans, a type of heparan sulfate proteoglycan, are conserved transmembrane domain proteins that also play a role in host cell signaling (433, 434). Several bacterial and viral pathogens have been shown to exploit binding to host cell surface SPGs to drive

117 signaling that promotes endocytic uptake to gain entry into the cell (430). One recent study indicated that T. gondii Type II strains can enter murine macrophages via phagocytosis, whereas Type I strains predominately enter host cells via active invasion (308). The parasite factors and host ligands contributing to this phenotype, however, have not yet been identified. It is possible that T. gondii SRS29C expression exploits SPG binding to gain entry into the cell via an endocytic pathway in a strain-specific manner to alter immune cell signaling as a means to ultimately regulate parasite proliferation.

Virulence is a complex, multi-factorial trait involving regulation of both host and pathogen factors. It is unlikely SPGs alone are solely responsible for the SRS29C- dependent phenotypes observed in this study. Several studies have reported diverse mechanisms by which SPGs can regulate inflammatory responses by interacting with modulators of the inflammatory responses, which include recruitment of FH to control complement activation and inflammation (121, 130, 391), as well as the binding of chemokines and cytokines to facilitate immune cell recruitment to sites of infection (435–

442). The structural basis for FH recruitment on host cells requires a joint dual recognition of SPGs and the C3d domain of C3b (130, 421). Because SRS29C was previously found to interact with C3b and FH and was further validated in this study using an SRS29C transgenic over expressing strain, we hypothesized that T. gondii SRS29C exploits this interaction to recruit FH. Thus, we engineered the transgenic SRS29C++ K16/17S strain to demonstrate a link between SRS29C-mediated FH recruitment and its ability to bind SPGs.

It is unclear from this study, however, how the parasite sequesters or retains SPGs on its surface. One conceivable scenario is that the parasite sequesters enzymatically cleaved

GAGs, which are released from the host cell surface via ectodomain shedding (443),

118 mediated by host metalloproteases upon infection or injury (444). Alternatively, soluble

SPGs are also abundantly present in serum (445), which could exist as a potential source of SPGs in our in vitro assays.

The complement system is a potent innate immune effector system that is stringently regulated by T. gondii to protect from complement attack (Chapter 2). The host also has several regulatory mechanisms in place to avoid uncontrolled inflammation and tissue damage. Host regulators achieve protection by inactivating complement activation product C3b to stop the cascade from amplifying and progressing. The importance of FH as a critical factor in regulating complement activation to control inflammation in the host is highlighted by the mutations and polymorphisms in the human FH gene that affect FH interactions with tissue specific SPGs and are associated with dysregulated complement responses that contribute to inflammatory diseases, tissue destruction, and autoimmunity

(96). Here, we showed that T. gondii exploits FH SPG-dependent recruitment to inactivate

C3b and down regulate complement activation. Western blot analyses had shown that there were intrinsic differences in the amount of iC3b between the SRS29C++ and SRS29C++

K16/17S strains, whereby abrogated SPG binding and FH recruitment resulted in less iC3b on the surface of SRS29C++ K16/17S parasites. Thus, another component to consider in the SRS29C-SPG interaction is downstream effects of FH recruitment in down regulating complement activation. Complement inactivation products have important effector functions in vivo that impact inflammatory processes, phagocytosis, clearing apoptotic cells, and stimulating adaptive immune responses, and maintaining host homeostasis (5).

Thus, alterations in the production of these products as a result of SRS-mediated complement regulation could alter several important effector functions.

119 Infection of C3 deficient mice demonstrated that the expression of SRS29C and C3 are both required for attenuating virulence, however the C3 effector function involved in this phenotype has not been resolved. In Chapter 2, we established Δsrs29C were equally resistant to serum killing despite reduced levels of FH and C4BP, suggesting complement- mediated lysis was not an effector function likely contributing to protection observed in these in vivo studies. Because there are several receptors for C3b and C3a, it is very likely that innate immune cells responding to these effector proteins could contribute to alterations in immune signaling and cytokine production. Increased complement deposition on SRS29C++ strains indicates that these strains are likely targeted for phagocytosis.

Conversely, in Chapter 2 we showed the loss of SRS29C as a C3 acceptor resulted reduced

C3b deposition and FH recruitment. iC3b is recognized by complement receptors CR3 and

CR4, which mediates phagocytic uptake of opsonized microbes and apoptotic cells.

Considering invasion is the major mode of parasite entry into host cells, studies evaluating the role of complement-mediated phagocytosis are lacking. One could speculate, however, that any parasite-driven alterations in the amount of these effectors being sensed by complement receptors on immune cells could contribute to changes in immune recognition, endocytosis, or host cell signaling. To determine if parasites exposed to serum could impact cytokine production, we infected BMDMs in vitro and observed dampened cytokine and chemokine production compared to control parasites. Thus, it is possible that iC3b-coated parasites are similarly impacting cytokine production in vivo. However, these studies could not determine if complement-mediated phagocytosis had occurred, as flow cytometric assays conducted here only stained for parasites and not invasion or phagocytic markers.

120 Further studies are required to determine if complement-mediated phagocytosis contributes to these changes in cytokine production.

SRS29C was previously shown to be covalently modified by C3b. C3b is also known to impact adaptive immune responses. The interaction of complement receptor 2

(CR2) and C3dg-covalently bound antigen has been shown to significantly lower the threshold for B cell activation and antibody production (137, 143). In fact, immune serum from mice infected with RH SRS29C++ parasites generate SRS29C specific antibodies whereas mice infected with wild type parasites did not (unpublished results), indicating that this interaction may facilitate the immunogenicity of SRS29C through its interactions with complement. However, how interactions with SPG and FH, and the resulting decrease in iC3b on SRS29C++ K16/17S impacts the antibody response has not been evaluated.

Further studies with C3 deficient mice would be required to confirm this is a C3-driven phenotype.

Anaphylatoxins C3a and C5a are complement effectors that also to play a role in modulating inflammatory responses. Traditionally, C3a and C5a are known as inflammatory molecules released upon C3 activation in response to invading pathogens.

Anaphylatoxins have multiple effector functions by interacting with C3a and C5a receptors

(C3aR, C5aR) on various immune and non-immune cells, including promoting inflammation, chemoattraction of immune cells to the site of infection and vasodilation.

By limiting the magnitude of alternative pathway complement activation, FH also regulates anaphylatoxin production. It is possible that the unregulated inflammatory responses observed in mice infected with Δsrs29C and SRS29C++ K16/17S strains could be attributed to changes in complement regulation on Δsrs29C and SRS29C++ K16/17S strains as a

121 result of reduced FH recruitment, resulting in increased levels of C3a and C5a released in the local environment that may impact recruitment of innate immune cells and cytokine responses. Lending support to this possibility, recent studies with bacterial pathogen

Streptococcus pneumoniae revealed that alternative pathway regulation by FH reduced C5a concentrations and decreased proinflammatory cytokine production in human peripheral blood mononuclear cells (446). Further, signaling through C5aR1 was shown to exacerbate immunopathology of Neisseria meningitis sepsis in mice (447). Further evaluations of anaphylatoxin production during acute infection with Δsrs29C and SRS29C++ K16/17S strains are necessary to consider if this is an important factor in pathogenesis.

In line with this rationale, C3 deficient animals, which lack C3a in serum, exhibited less IFN-g, IL-6 and IL-18 systemically compared to WT mice, suggesting the mice failed to induce the appropriate Th1 response. However, there were no significant differences in parasite burden between wild type and C3-/- mice, suggesting that despite these reductions in cytokine production, they were both able to control parasite proliferation, thus raising the question of what contributes to mortality in these mice. Furthermore, this rationale does not explain the increased production of inflammatory cytokines by splenocytes from C3-/- infected animals. The recent discovery of intracellularly generated complement and secreted products showed that C3a and C5a can induce signaling through their cognate receptors to modulate Th1 responses (158), demonstrating serum-derived and locally produced complement by immune cells have distinct functions (448). Despite C3 deficiency, we observed similar levels of C5a in the serum of these mice, indicating a C3 bypass mechanism that activates C5. Furthermore, C5a is also known to be generated by phagocytic cells, suggesting an additional serum-independent source of C5a (449). C5a is

122 reported to be more potent than C3a (450), thus the increase in cytokine production by C3 deficient splenocytes could possibly be attributed to compensatory C5a production in the absence of C3a, which may induce a more potent Th1 response. Thus, it is not clear from these studies if the virulence phenotype we are observing in C3 deficient mice is due to loss of a systemic C3 response, or a dysregulation of the complement response due to an unanticipated bypass in C5 activation.

Modulation of pro-inflammatory Th1 response, which includes IL-12 and IFN-γ production, is critical for regulating T. gondii proliferation and parasite persistence. In this chapter, we establish that the complex interactions of T. gondii SRS29C with extracellular host factors is a critical event during the establishment of infection that modulates downstream immune signaling and impacts pathogenesis. Here, we propose a novel strategy by which T. gondii SRS29C exploits complement regulation to promote parasite persistence and host survival by modulating inflammatory Th1 responses. We showed that

SRS29C binds host SPGs to facilitate recruit of soluble regulator Factor H to control complement activation, a known mediator of inflammation. Furthermore, we demonstrated that C3 and SRS29C-mediated SPG binding alter pathogenesis and inflammatory cytokine production in vivo that was not due to changes in parasite burden early in infection, but as a result of alterations in immune priming. Taken together, these findings have assigned the first biological effector function to a T. gondii SRS protein in regulating acute virulence, thus significantly expanding our understanding of the host ligands that contribute to the diverse biological functions of the Toxoplasma SRS superfamily of proteins and ultimately how these interactions promote parasite persistence.

123 3.4 Materials and Methods

3.4.1 Construct engineering, protein production and purification

The tandem SRS domains of SRS29C (Pro55 to Gly311) from Type I (GT1; SRS29C-I) and Type II (ME49; SRS29C-II) T. gondii were amplified from genomic DNA and cloned into a modified pAcGP67B vector (Pharmingen) with N and C-terminal thrombin cleavage sites in addition to a C-terminal hexahistidine tag. The tandem SRS domains of SRS29B and SRS16C were cloned as previously described (422). SRS homodimers were generated by subcloning the tandem SRS domains into a pAcGP67B vector containing a C-terminal leucine zipper (modified from (451)) between the thrombin cleavage and hexahistidine sequences. SRS29C (II) mutants were subsequently generated by site-directed mutagenesis using the QuikChange XL kit (Stratagene) with the dimer clone as a template.

For expression in insect cells, each clone was transfected with linearized

Baculovirus into Sf9 cells and amplified to a high titer. Hi-5 cells at 1.8x106 cells/mL were infected with amplified virus for 65 hours, after which time the supernatant was harvested, concentrated by tangential flow and applied to a HisTrapFF Ni affinity column (GE

Healthcare). Elution fractions were analyzed by SDS-PAGE and pooled based on purity.

For crystallization, the hexahistidine tag and/or dimerization domain was removed by thrombin and the protein further purified by gel filtration (Superdex 16/60 200) and anion exchange chromatography (Source 30Q). Purified wild-type and mutant SRS29C were concentrated to between 28 and 36 mg/mL in HEPES buffered saline (HBS: 20mM HEPES pH 7.5, 150mM NaCl).

124 3.4.2 Crystallization and data collection

Crystals of SRS29C (I and II) were initially identified in the Index screen (Hampton

Research) and refined to a final condition of 12% PEG3350, 0.1 M HEPES pH 8.0, 5 mM cobalt chloride hexahydrate, 5 mM nickel chloride hexahydrate, 5 mM cadmium chloride monohydrate, 5 mM magnesium chloride hexahydrate. For both proteins, crystals were observed after 2 days and grew to a maximum size of 0.5 x 0.4 x 0.2 mm after 4 days at

293 K. The final drops consisted of 1.0 μL protein at 36 mg/mL and 1.0 μL reservoir solution and were equilibrated against 100 μL of reservoir solution. Diffraction quality was increased by streak seeding into drops set with protein at 28 mg/mL. Cryoprotection of the

SRS29C crystals was achieved by soaking in mother liquor supplemented with 12.5% glycerol and flash cooling at 100K directly in the cryo stream. Diffraction data were collected on beamline 9-2 at SSRL (Stanford Synchrotron Radiation Lightsource).

3.4.3 Data processing, structure solution and refinement

Diffraction data were processed using iMosflm (452) and Aimless (453) in the CCP4 suite of programs (454). For SRS29C (I), initial phases were obtained by molecular replacement

(MR) using PHASER with the individual D1 and D2 domains of a hybrid model of

SRS29B (265) and SRS16C (422) pruned by chainsaw (455). For SRS29C (II) and the mutants, initial phases were obtained by MR using PHASER with the finalized SRS29C

(I) model. Manual building and selection of solvent molecules was performed using COOT

(456) and refinement steps carried out using Phenix.refine (457). Complete structural validation was performed in MolProbity (458). The Ramachandran plot for each structure shows excellent stereochemistry with greater than 97% of the residues in favored conformations and no residues modeled in disallowed orientations. Overall 5% of the

125 reflections were set aside for calculation of Rfree. Data collection and refinement statistics are presented in Table 1.

3.4.4 Sulfated proteoglycan binding studies

For the pulldown assays, a slurry of heparin-agarose beads (Sigma) (30 μL) was pre- equilibrated in binding buffer (20 mM HEPES, 50 mM NaCl, pH 7.5) and added to 10 μg of each protein to a final reaction volume of 600 μL. The mixture was then incubated at room temperature for 45 min with inverting every 10 min and subsequent centrifugation at

1,600 rpm for 5 min. The supernatant was removed, and the beads were washed 3 times with cold binding buffer. All buffer was removed, and 15 μL of 4X SDS loading buffer was added to the beads. Samples were heated to 95 °C for 5 min, centrifuged at 1,600 rpm for 5 min, and separated on a 15% SDS polyacrylamide gel.

For native gel shift assays, purified proteins (4 μg) in HBS were incubated for 30 minutes at room temperature with heparin sodium salt (Sigma), de-N-sulfated heparin (V- labs), or heparan sulfate (Sigma) in a 1:2, 1:5, or 1:10 protein to heparin molar ratio.

Solutions were run on 8-25% gradient gels using native buffer strips and the PhastGel system (GE Healthcare).

3.4.5 Parasite culture

Toxoplasma gondii tachyzoites were maintained by in vitro passaging in confluent monolayers of human foreskin fibroblasts (HFF) in Dulbecco modified Eagle medium

(DMEM) with 10% fetal bovine serum (FBS), 2mM glutamine, penicillin, streptomycin and 10µg/ml gentamicin at 37°C, 5% CO2. Parasites were counted using a hemocytometer.

126 3.4.6 Plasmid constructs and parasite strains

The extracellular domain of the coding region of the SRS29C gene was amplified from

Type I RH and Type II 76K strain genomic DNA using the sense primer, 5’-

CGGGATCCGCCGTACAGATACGAGC-3’ and the antisense primer, 5’-

AAAACTGCAGCACTCCCTCCACCCCCCTG-3’. The resulting PCR fragments were cloned into a BamH-I and Pst- I cut SRS29B pBluescript-based expression plasmid,

Pru1.6Ava (Grigg, unpublished data) that replaces the extracellular domain of SRS29B with SRS29C to produce the Pru1.6Ava SRS2-I and –II plasmids. The Pru1.6Ava-SRS2-I and SRS2-II plasmids contain the extracellular domain of the coding sequence flanked by sequences corresponding to the SRS29B promoter, un-translated regions, signal peptide, and GPI anchor. These plasmids were transfected into the parental parasite RH∆ strain for this study (WT). WT pertains to a Type I RH strain that has a disrupted hypoxanthine- xanthine-guanine phosphoribosyl transferase (HXGPRT). The pHXGPRT plasmid contains the HXGPRT open reading frame flanked with the regulatory regions isolated from the Di-Hydro Folate Reductase (DHFR) gene. The pGFP-LUC plasmid contains separate cassettes for GFP (driven by the GRA4 promoter) and LUC (driven by the DHFR promoter).

3.4.7 Attachment/invasion assays

For the plaque assay, 1000 freshly lysed tachyzoites were allowed to invade HFF cells for the 15 min, 30 min, 1hr and 2hr time points and 200 for the 24hrs time point. Extracellular parasites were then washed off four times with PBS. Plaques were counted after ~6 days.

For the bioluminescence-based attachment/invasion assay, freshly lysed luciferase expressing clones and the parent cell line, 0.5 million each, were inoculated into 24-well

127 plates containing human fibroblast (HFF) cells. Parasites were allowed to invade HFF cells for 15 minutes, 45 minutes, 2 hrs, 24 hrs, or 48 hrs respectively, after which extracellular parasites were washed off four times with PBS. 50 µg of luciferin substrate dissolved in

PBS was added to each well and the plates were imaged to detect the total flux of photons per well using Xenogen’s in vivo imaging system (IVIS).

3.4.8 Flow cytometry to measure C3b deposition and FH recruitment

++ complement activating buffer HBSS (Hanks Buffered Saline Solution, 1mM MgCl2,

0.25mM CaCl2). All serum was tested by IFA and flow cytometry for the presence or absence of human anti-Toxoplasma IgG prior to use. Complement activation was stopped using cold PBS and washed two times with cold PBS to remove excess serum. Parasites were then fixed with 1% PFA solution for 10 minutes, washed, and stained with mouse α- human C3b/iC3b antibody (1:500) (Cedarlane), or goat α -human Factor H (1:2000)

(CompTech), followed by α-mouse APC (eBiosciences) or α-goat APC conjugated antibody (Jackson Laboratories), both at 1:1,000. 20,000 events were collected on a BD

Fortessa flow cytometer and analyzed by FACs DIVA, FlowJo and Prism software.

3.4.9 Mouse infections

Eight-week-old C57BL6 (Jackson Laboratories) mice and C3-/- (Jackson laboratories) were infected intraperitoneally (i.p.) with tachyzoites. Parasites cultured in HFFs cells were washed in twice in PBS by centrifugation at 1250 rpm for 10 minutes and counted on a hemocytometer. Mice were injected with 2x103 tachyzoites in 500μl in incomplete RPMI

128 for infections with CZ1 Type II strains or 25 tachyzoites in 500μl in incomplete RPMI for infections with RH Type I strains.

3.4.10 Mouse infection and bioluminescent in vivo imaging

7-14-week-old female CD1 outbred mice were infected by intraperitoneal injection with a total of 50 parasites diluted in 200-400 µl of PBS. Xenogen’s in vivo imaging system (IVIS) was used for data acquisition and analysis. D-luciferin, dissolved in PBS at 15.4 mg/ml and filtered through a 0.22 µm filter, was used as the substrate for firefly luciferase. Mice were injected with 200 µl of an equal mixture of the luciferin and PBS (3 mg). Mice were maintained for 5-10 minutes to allow sufficient circulation of luciferin and subsequently anesthetized for 3-5 minutes with haloethane. Exposure time during image acquisition ranged between 10 seconds to 3 minutes. Minimum photons were consistently maintained for all images. Pseudocolor representations of light intensity were overlaid onto grayscale images of the mice with red being the highest and blue being the lowest intensity.

3.4.11 Plaque assays to quantify parasite load

Mice were euthanized at day 4 post infection when infected with RH strains or day 6 post infection when infected with CZ1 strains. Peritoneal fluid was collected by injecting 5ml of sterile PBS into the peritoneal cavity of euthanized mice. Spleens and lungs were harvested and homogenized over 70-μm strainers in 5ml of X-VIVO-20 serum free media.

Ten-fold serial dilutions starting with 500 μl of resuspended homogenate were added to

HFF monolayers in 12 well plates and cultured undisturbed in DMEM media for 7 days.

HFFs were fixed in methanol and stained with crystal violet. Plaques were counted using an inverted microscope and parasite loads were expressed as plaque forming units (PFU) per organ.

129 3.4.12 Ex vivo splenocyte cultures

Spleens were homogenized over 70-μm strainers in 5 ml of X-VIVO-20 serum free media.

After red blood cell lysis (Lonza ACK lysing buffer), splenocytes were plated in a 96-well flat bottom plate at 5x105 splenocytes per well in 250 μl of X-VIVO serum free media and cultured for 24 hours. 250 μl of supernatants were collected for analysis.

3.4.13 Measuring cytokine production from serum and ex vivo splenocyte cultures by

ELISA

Serum was prepared from blood collected by cardiac puncture. Blood was allowed to clot for 30 minutes and then spun at 1000 rpm for 15 minutes to extract serum. Supernatants of ex vivo splenocyte cultures were collected as previously described. Samples were analyzed using ELISA kits for IL-12p40, IFN-γ, TNF-α, IL-6, IL-18 and IL-10 according to manufacturer’s instructions (R&D Systems).

3.4.14 BMDM culture

Bone marrow from of C57BL6 mice were harvested from femurs and hind leg bones. Cells were washed with RPMI medium and resuspended in RPMI medium with 10% FBS, 10

U/mL penicillin, and 10 μg/mL streptomycin (Gibco). Macrophages were differentiated with 30% L929 conditioned medium was added to RPMI medium with 10% FBS. The cells were cultured in 100 × 20 mm dish plates (Costar; Corning Inc., Corning, NY), supplemented media at day 4. At day 7, macrophages were then harvested and plated into

96-well plates at 5 × 105 cells/well for T. gondii infections.

3.4.15 In vitro infection of BMDMs

Cultured parasites from a freshly lyses HFF monolayer were collected, filtered, and washed twice. Parasite were resuspended in RPMI media and counted. Half of the parasites

130 were first opsonized by incubated in 40% non-immune human serum (NHS) for 20 minutes at 37°C and washed twice with cold RPMI media. Cells were infected at MOI of 3 and incubated for 20 hours 37°C. Supernatants were collected, and macrophages were then analyzed for infectivity by flow cytometry.

3.4.16 Luminex assay

Supernatants from BMDM experiments were collected at 20 hours and analyzed by

Luminex (Millipore) according to the manufacturer’s instructions.

3.4.17 Measurement of C3a and C5a levels in mouse serum:

A 96-well plate was coated with purified 2 μg/ml of rat anti–mouse C3a or rat anti-mouse

C5a antibody in 0.1 M NaHPO4 coating buffer, pH 9.5 (BD Biosciences), overnight at 4°C.

The plate was washed three times with PBS containing 0.05% Tween 20 and blocked with

PBS/10% FBS for 1 h at room temperature. Samples were diluted 1:100 in blocking buffer and incubated for 2 h, followed by five washes. Bound C3a and C5a was detected using biotinylated 1 μg/ml of rat anti–mouse C3a or C5a antibody and avidin–horseradish peroxidase. Reactive C5a levels were measured using 2,2′-Azino-bis (3- ethylbenzothiazoline-6-sulfonic acid) (Sigma-Aldrich) for 30 min, and the reaction was stopped using 2 N H2SO4. The plate was read at an OD of 405 nm.

3.4.18 Statistical analysis

Data were expressed as mean ± standard error of the mean of at least two independent experiments. Statistical analyses were performed using Student’s t test between wild type strains and transgenic or knock out parasites, and mean differences, which are denoted by an asterisk, were considered statistically significant if *p<0.05, **p<0.001, ***p<0.0001.

131 3.5 Figures

Figure 3.1. Tandem SRS domains of SRS29C reveal an apical basic surface. (A) Domain organization of SRS29C. Polymorphisms between Type I and Type II SRS29C: 113, N/S; 271, R/M; 295, E/D; 338, S/G. SP, signal peptide; GPI, predicted glycophosphatidylinositol anchor site. Disulfide connectivity is shown as black connectors beneath each domain. (B) Cartoon backbone representation of SRS29C in the predicted orientation to the parasite surface. D1 and D2 are colored green, with the interdomain linker colored dark grey. Disulfide bonds shown as ball-and-stick. Polymorphic sites are colored orange. (C) Apical view of an electrostatic surface of SRS29C D1 shows an extended basic region comprised of five Lys and one Arg residue. (Crystal structure was solved by the Martin Boulanger group at the University of Victoria).

132 Figure 3.2. SRS29C dimer dependent binding to heparin localized to basic groove. (A) Predicted orientation of the GPI-anchored SRS29B homodimer on the parasite surface, and a model of the C-terminal leucine zipper based homodimerization domain (represented by PDB ID: 2ZTA) that enables dimerization in solution. (B) Overlay of size exclusion traces of SRS29C monomer and dimer, showing homodimerization of the engineered form of SRS29C. Globular protein standards: I, 75 kDa; II, 43 kDa. Inset – SDS PAGE analysis of the purified proteins with reductant shows expected migration. (C) Pull down assays with heparin agarose beads. S – protein remaining in solution; PD – protein pulled down. The arrow in the far-right panel indicates a substantial portion of SRS29C dimer pulled down with heparin compared to other SRS monomers and dimers. (Gel shift performed by Michelle Tonkin, University of Victoria) (D) Native gel analysis of SRS29C dimer incubated with heparin in 1:2, 1:5, or 1:10 molar ratios show an increasing upshift with increasing heparin concentration, while the SRS29C monomer incubated with heparin in a 1:10 molar ratio shows only a very minor blurring of the protein band. (E) Native gel analysis of SRS29C alone or pre-incubated with heparin, de-N-sulfated heparin, or heparan sulfate confirms the dependency on sulfation of the heparin. (F) Residues chosen for site- directed mutagenesis mapped onto the modeled dimer of SRS29C. (G) Native gel analysis of specific SRS29C dimer mutants shows a loss of heparin binding by mutants within the predicted basic groove.

133 Figure 3.3. SRS29C-medaited FH recruitment is SPG-dependent. 1x106 RH (black square), RH SRS29C++ (black triangle) and RH SRS29C++ K16/17S parasites were incubated in 10% NHS over 60 minutes at 37°C and strained with (A) goat-anti-human FH (1:2,000) or (B) mouse anti-human C3b (1:500) and analyzed by flow cytometry. RH parasites in 10% heat inactivate serum (D56, open square, dotted line) were used a negative control (C) and (D) represents an average of three independent experiments done in duplicate at the 30-minute time point. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, *p<0.05. (E) Western blot analysis of the form of C3b on RH, RH SRS29C++ and RH SRS29C++ K16/17S parasites incubated in 10% NHS for 20’. The transgenic strains used in these assays were generated by Dr. Viviana Pszenny, NIH.

134 Figure 3.4. SRS29C trans-overexpression limits wild type parasite proliferation in vivo. (A) Attachment and invasion assays in HFF and J774 cells at different time points (B) Quantification of bioluminescence in (A) represented as total flux (photos/sec) (C) CD-1 outbred mice were infected intraperitoneally with 50 GFP-Luc positive wild type tachyzoites alone, or with a mix of 50 GFP-Luc positive wild type parasites + 50 “dark” (GFP-Luc negative) RH SRS29C++ parasites. Parasite burden was measured via bioluminescent detection in photons/sec/cm2 at days 3, 5, and 7 post infection.

135

Figure 3.5. SRS29C over expression alters pathogenesis in SPG-dependent manner. (A) Survival of CD-1 outbred mice were infected with 25 tachyzoites of RH (red), RH SRS29C++ (green) or SRS29C++ K16/17S (blue) parasites via footpad injection. (B) Parasite burden was measured by bioluminescent detection in photons/sec/cm2 at day 9 post infection. Transgenic lines used in this figure were generated by Dr. Viviana Pszenny (NIH).

136 Figure 3.6. SRS29C-SPG interaction alters early cytokine responses. Mice were infected with 25 tachyzoites intraperitoneally with RH (black bar), RH SRS29C++ (gray hatched bar), and RH SRS29C++ K16/17S strains (open bar). Mice were sacrificed 4 days post infection. Parasite burdens were determined by counting plaque forming units (PFUs) by plating peritoneal fluid (A) and homogenized spleens (B) onto HFF monolayers in 12 well plates. Serum was collected at day 4 post infection and evaluated for IL-18 (C) and IL-12 and IFN-g (D) by ELISA. Data are an aggregate of two independent experiments, n=5. Data are represented as mean ± SEM and p-values were obtained using Student’s t- test, *p<0.05, **p<0.01.

Figure 3.7. SRS29C-mediated attenuation of virulence is C3-dependent. (A) Survival of 6-8-week-old C57BL6 female mice infected with 2x103 WT CZ1 (black) and CZ1Dsrs29C tachyzoites (red) intraperitoneally. Data is representative of two independent experiments, n=11. (B) Survival of C3-/- mice infected with 2x103wild type (black dotted line) and Dsrs29C tachyzoites intraperitoneally (red dotted line), n=3.

137 Figure 3.8. SRS29C expression alters inflammatory cytokine production in vivo. 6-8- week-old C57BL6 female mice infected with 2x103 wild type CZ1 (gray bar) and Dsrs29C tachyzoites (open bar) intraperitoneally, or PBS alone (naïve, black bar). Mice were scarified 6 days post infection (A) Parasite burden was assessed by counting plaque forming units (PFUs) of peritoneal fluid and homogenized spleens and lungs plated onto HFF monolayers in 12-well plates. Serum was collected to measure systemic IL-12 (B) and IFN-g (C) levels. Ex vivo splenocytes were cultured for 24 hours and supernatants were collected to measure local IL-12 (D), IFN-g (E), TNF-a (F) production. Data are an aggregate of two independent experiments, n=4. Data are represented as mean ± SEM and p-values were obtained using Student’s t-test, **p<0.01, ***p<0.001.

138 Figure 3.9. C3 alters inflammatory cytokine production during acute T. gondii infection. 6-8-week-old C57BL female mice (black bar) or C3-/- female mice (open bar) infected with 2x103 wild type CZ1 parasites or PBS alone (naïve, gray hatched bar) intraperitoneally. Mice were sacrificed 6 days post infection. Parasite burden was assessed by counting plaque forming units (PFUs) of peritoneal fluid (A) and homogenized spleen (B) plated onto HFF monolayers in 12-well plates. Serum was collected to measure systemic IL-12 (C), IFN-g (D), IL-6 (E), and IL-18 (F) levels. Ex vivo splenocytes were cultured for 24 hours and supernatants were collected to measure local IL-12 (G), IFN-g (H), and IL-6 (I) production. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, *p<0.05, **p<0.01.

139

Supplemental Figure 3.1. SRS29C regulates parasite burdens and inflammation in the brains of infected mice. (A) Survival of 6-8-week-old C57BL6 mice infected with Type II strain Prugniaud (Pru) strain. (B) Brain cyst burdens. (C) Histological H&E staining on brain sections from WT infected and Dsrs29C infected mice shows increased inflammation of the meninges (upper right panel), infiltrate of inflammatory mononuclear cells (middle right panel) and the presence of tachyzoites (bottom right panel) in the brains of mice infected with Δsrs29C parasites. Histopathology was done by Dr. Alessandra Commodaro, NIH.

140 Supplemental Figure 3.2. Serum levels of C3a and C5a in C3-/- mice. Serum levels of C3a (A) and C5a (B) at 4, 6, and 8 days post infection from mice infected with 40 cysts Type II ME49 parasites intraperitoneally. C3a and C5a ELISAs were performed by Dr. Alessandra Commodaro, NIH

Supplemental Figure 3.3. BMDM infection with serum coated parasites dampens cytokine and chemokine production in bone marrow derived macrophages. BMDMs were infected with parasites with and without prior incubation in 40% NHS for 30 minutes at 37°C at MOI of 3. (A) Supernatants were collected after 20 hours and cytokine and chemokine production were measured using a Luminex assay Macrophages were stained for intracellular T. gondii and measured by flow cytometry. (B) Infectivity is expressed as percent of cells positive for T. gondii. Data is an aggregate of two independent experiments. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, *p<0.05, **p<0.01.

141 CHAPTER 4: TOXOPLASMA SURFACE COAT PROTEIN SRS57 IS

COVALENTLY MODIFIED BY COMPLEMENT AND REGULATES PARASITE

PATHOGENESIS IN MOUSE MODEL OF ACUTE INFECTION3

4.1 Introduction

The protozoan parasite Toxoplasma gondii is a widespread intracellular pathogen that is the etiological agent of toxoplasmosis, causing significant morbidity and mortality in humans and animals worldwide (219). A critical factor in T. gondii pathogenesis is the ability of the parasite to disseminate to various tissues and establish a chronic infection.

The capacity of T. gondii to cross various biological barriers and infect many cell types underlies the disease that accompanies toxoplasmosis (459), which includes crossing the placenta to cause congenital infection, or spread to the eye in healthy adults. As a consequence, it is the leading cause of infectious uveitis worldwide (215, 226, 244).

Toxoplasmosis can be particularly serious in chronically infected immunocompromised individuals who are at risk for reactivated disease (221). In the absence of adequate immunity, uncontrolled parasite proliferation during reactivated disease can be fatal. In healthy individuals, sufficient immunity is able to control parasite replication, but it is unable to clear the chronic phase. To achieve this, T. gondii employs several strategies of immune evasion to permit its dissemination to various tissues, while concurrently

______3Citation: Sikorski, P.M., Pszenny, V., Raghavendran, R., Parker, M.L., Adedoyin, G., Boulanger, M.J., and Grigg M.E. Toxoplasma surface coat protein SRS57 is covalently modified by complement and regulates parasite pathogenesis in mouse model of acute infection. In preparation.

142 stimulating sufficient immunity to regulate parasite proliferation and support stage conversion. The Toxoplasma surface coat proteins are postulated to play a role in regulating host immunity and establishing persistent infection. The surface of Toxoplasma is populated by a large superfamily of developmentally regulated surface proteins, known as the SRS (SAG1-related sequences) proteins (259).

SRS proteins are thought to play a critical role in promoting parasite attachment to host cells as well as immune evasion to facilitate parasite dissemination and persistence.

Early in vitro studies attributed attachment and adhesion to several T. gondii SRS proteins, including SRS29B (formerly SAG1) and SRS57 (formerly SAG3) (258, 384). Moreover, blocking SRS29B or SRS34 (formerly SAG2) obstructed host cell invasion (262, 263), suggesting that these proteins are critical in promoting infectivity. As a generalist parasite, the remarkable ability of T. gondii to infect almost any cell type in a broad range of hosts implies that attachment relies on the ability of the parasite to recognize conserved and widely distributed surface molecules or constituents of extracellular matrices. Many pathogens, including bacteria, viruses and parasites, mediate attachment to host cells by binding glycosaminoglycan chains of proteoglycans (460–462), which are a major component found ubiquitously on all mammalian tissues.

Several studies have shown that T. gondii possesses a lectin-like activity that is associated with their ability to interact with host sulfated proteoglycans (SPGs) (255–257,

463). Further, cells lacking proteoglycans or sulfation were more resistant to T. gondii infection (256, 258). SRS29B and SRS57 have previously been shown to bind SPGs with strong affinity (258, 264). Whereas the SRS29B-deficient mutant shows no appreciable deficit in host cell infection (262), the SRS57 deficient parasites were significantly

143 impaired in host cell attachment and murine infection competency (464). Follow-up binding studies have demonstrated a specific interaction between SRS57 and heparin (258).

The crystal structure of SRS29B rationalized the structural basis for these observations, by revealing a dimer dependent basic groove that was ideally suited to coordinate a negatively charged cellular ligand, such as heparan sulfate (265). Based on this structure, the model developed for SRS57 identified a substantially more basic groove consistent with its role as a major surface adhesin capable of recognizing negatively charged SPGs (265).

Evasion of the complement system is also a critical step in the establishment of infection. This important innate immune response limits pathogen dissemination by several means, including targeting pathogens for elimination by direct lysis or phagocytosis, stimulating B cells (144), and regulating T cell differentiation (152, 158). Previous work has demonstrated that the surface of T. gondii tachyzoites is covalently modified by complement proteins upon exposure to non-immune human serum (173). In Chapter 2, it was determined that SRS29C is a nucleophilic target of C3b, however, its expression did not account for all C3b binding. The molecular basis for T. gondii resistance to complement-mediated killing was attributed to a cooperative function of SRS29C

(formerly SRS2) and SRS25 in recruiting host regulators Factor H and C4b-binding protein to down regulate complement activation on the parasite surface. However, Factor H- mediated regulation of the alternative pathway was shown to significantly contribute to serum resistance.

Factor H, an important negative regulator of the alternative pathway, is a single chain glycoprotein consisting of 20 homologous domains known as short consensus repeats

(SCRs) or complement control proteins (CCPs). Factor H typically identifies and binds

144 host cell surfaces through the recognition of C3b and host “self” molecules, which include sialic acid and SPGs (51). Factor H has several C3b and SPG binding sites (121, 124, 127,

128, 130, 420, 421). Several studies have determined that the structural basis for Factor H recruitment to host cells is dependent on its ability to simultaneously bind both C3b and host SPGs (130). Because T. gondii binds both C3b and SPGs, we postulated that the capacity of SRS proteins to recruit Factor H was dependent on their capacity to bind SPGs and C3b. In Chapter 3, we demonstrated that SRS29C-mediated FH recruitment was dependent on its ability to bind SPGs.

The purpose of this study was to determine if the predicted capacity of SRS57 to bind SPGs was likewise a mechanism for SRS57-mediated Factor H recruitment to the parasite surface. To answer these questions, ∆srs57 parasites were generated to test the role of SRS57 in binding C3b and FH. SRS57 was identified as an additional target for C3b in vitro. Although ∆srs57 parasites also had a reduced capacity to recruit FH, the crystal structure determined it did not support SPG binding, indicating T. gondii could employ

SPG-independent mechanisms for FH recruitment. However, we identified several immunogenic tachyzoite proteins that bound heparin sulfate with high affinity. In vivo studies indicated that like SRS29C, the interaction between SRS57 and C3b plays an important role in limiting pathogenesis, thus promoting parasite persistence in mice.

4.2 Results

4.2.1 Targeted deletion of SRS57 locus in Type I RH and Type II CZ1 strains

In order to study the role of SRS57 in complement activation, ∆srs57 deficient strains were generated in the RH Type I and CZ1 Type II genetic background. The genomic locus of SRS57 was first disrupted by homologous recombination using an HXGPRT mini gene

145 as the positive selectable marker in the RH∆hxgprt strain (Figure 4.1A). Screening for independent clones deficient in SRS57 expression from three separate transfections was performed by flow cytometry using an anti-SRS57 monoclonal antibody (1F12, kindly provided by Dr. Stan Tomavo). A total of 10 ∆srs57 clones were obtained. Parasite lysates from these clones were treated with GPI-PLC to expose the cross-reacting determinant

(CRD) present in surface-associated SRS proteins that possess a GPI-anchor, GPI-PLC treated lysates were subjected to SDS-PAGE, transferred to nitrocellulose and probed with anti-CRD antibodies. Consistent with the result of the flow cytometry analysis, a band corresponding to SRS57 was absent in the three ∆srs57 clones (Figure 4.1B- lanes 3, 4 and 5). Moreover, there did not appear to be any compensatory change in the surface expression of other GPI-linked SRS surface coat proteins (Figure 4.1B). In order to restore the expression of SRS57 at a heterologous site within the T. gondii genome, and to generate transgenic parasites expressing the green fluorescence protein (GFP) and firefly luciferase protein (LUC) for in vivo imaging experiments, ∆srs57 parasites (as well as the previously published French RH∆srs57 mutant CL55, kindly provided by Dr. S. Tomavo) were co- transfected with a 6.7kb PCR amplified genomic locus of SRS57, encompassing the 5’ and

3’ UTRs and the promoter region as well as a pGFP-LUC plasmid, containing cDNA sequences of GFP and LUC. The strategy for complementation is depicted in

Supplemental Figure 4.1. After transfection, parasites positive for GFP were sorted by flow cytometry and GFP positive clones were assessed for surface expression of SRS57.

No difference was detected in the expression of SRS57 between wild type (WT) parent and the complemented clones by flow cytometry (Figure 4.1C- left). The expression of

146 SRS34A, a highly immunogenic surface antigen, was unchanged in the knockout and complemented parasites (Figure 4.1C (right)).

To generate a ∆srs57 clone in the CZ1 Type II strain, we took a CRISPR/Cas approach to disrupt the SRS57 locus by inserting the HXGPRT mini gene after the SRS57 signal peptide in a recently generated CZ1∆ku80∆hxgprt. A single guide RNA was designed to introduce a break within the 5’ end of the SRS57 locus. The HXGPRT gene was amplified using primers that possessed 40bp of homology with the SRS57 gene either side of the guide RNA cut site (Figure 4.1D). After transfection and drug selection, screening of independent clones deficient in SRS57 was done by PCR. Clones positive for an HXGPRT at the SRS57 locus were then confirmed by staining for SRS57 using the anti-SRS57 monoclonal antibody (Figure 4.1E) and by Western blot (Figure 4.1F).

4.2.2 SRS57 expression impacts C3b and FH binding in vitro

Previous work demonstrated that T. gondii encodes several surface factors that are targets for complement effector protein C3b (173). Studies in Chapter 2 established that

SRS29C is one surface coat protein that is covalently opsonized by C3b. However,

Δsrs29C parasites did not completely abolish C3b deposition, indicating that at least one additional parasite factor was covalently modified by C3b. Our study also showed that opsonized parasites recruited both Factor H (FH) and C4b-binding protein (C4BP) to inactivate C3b to promote serum resistance (Chapter 2). Because FH was responsible for the majority of the resistance to serum killing, and FH recruitment to the surface of host cells is stabilized by binding both C3b and SPGs in close proximity (130), we postulated that SRS proteins recruit FH in an SPG- and C3b-dependent manner (Chapter 3). SRS57 has been previously shown to bind SPGs with high specificity (258) and a homology-

147 modeled structure predicted a binding groove capable of interacting with SPGs (265).

Hence, we took a reverse genetics approach to determine whether SRS57 is covalently modified by complement and is capable of recruiting FH.

Using flow cytometry, a time course study to analyze C3b deposition on ∆srs57 compared to WT parasites incubated in the presence of 10% non-immune human serum

(NHS) revealed that both Type I and Type II Δsrs57 parasites exhibited a significant

(~50%) reduction in C3b deposition (Figures 4.2A, 4.2B), which was also confirmed by western blot (Figure 4.2E).This suggested that, besides SRS29C, SRS57 is the other C3- acceptor on the surface of Toxoplasma tachyzoites. Next, Factor H recruitment was measured by flow cytometry. Δsrs57 parasites phenocopied the ∆56 NHS control indicating that in the RH background, tachyzoites did not accumulate Factor H on their surface (Figure 4.2C), whereas in Type II CZ1 strains there was a partial reduction in recruitment (Figure 4.2D), suggesting that SRS57’s predicted role in binding SPGs might play a more relevant role for Type I strains that only weakly express SRS29C.

4.2.3 SRS57 crystal structure does not support a role for binding SPGs

To determine if SRS57 also binds SPGs, its structure was determined using X-ray crystallography. SRS57 (Figure 4.3A) was produced recombinantly in insect cells and purified to homogeneity by nickel affinity and size exclusion chromatography (SEC). The protein eluted as a monomer from SEC and migrated as a doublet on SDS-PAGE due to differential N-linked glycosylation. TgSRS57 crystallized in space group P212121 with one molecule in the asymmetric unit and the final structure refined to a resolution of 1.59

Å. SRS57 was modeled from Tyr57 to Pro330, with an additional five residues at the C- terminus derived from the expression construct, and two disordered residues in an apical

148 loop (Leu102 and Gly103). Overall, SRS57 adopted a dumb-bell shaped architecture with

2 disulfide bond stabilized β sandwich domains connected by a proline rich linker (Figure

4.3B, inset). The D1 domain extended from Tyr57 to Glu198 and formed a five on four β sandwich, while D2 extended from Met204 to Val335 and formed a four on three β sandwich.

Comparison of the individual D1 and D2 domains of SRS57 to previously characterized members of the SAG1 family, SRS29B and SRS16C (BSR4), revealed that, in contrast to the membrane proximal D2 domains, the membrane distal D1 domains displayed increased structural variability (SRS57: SRS29B - 2.2 Å rmsd over 130 Cα atoms for D1 and 1.5 Å over 123 Cα atoms for D2; SRS57: SRS16C – 2.6 Å rmsd over 121 Cα atoms for D1 and 2.0 Å over 135 Cα atoms for D2). Similar to SRS16C, SRS57 incorporated an additional N- terminal β- strand (Figure 4.3C, indicated by red arrows) and extended β- strands in the top leaf of D1 compared to SRS29B. The two longest strands in the SRS57 D1 top leaf are of particular note, as they formed an extended and curved β- hairpin, where the analogous strands in both SRS29B and SRS16C are much shorter or barely recognized (Figure 4.3C, indicated by black arrows. These differences are particularly interesting since D1 is apically positioned and thus most likely to engage partners on the host cell surface. Further, the variability in the apical surface may affect dimerization. Homodimerization of SRS29B is necessary to form the basic groove that is predicted to coordinate a negatively charged ligand, possibly SPGs (265, 422, 465).

Notably, there was no evidence for dimerization either from the SEC or crystal packing of

SRS57. In fact, attempts to model the SRS57 dimer using the SRS29B scaffold were unsuccessful due to a series of steric clashes between the substantially more extensive β

149 sheet structure of SRS57 D1 (Figure 4.3C- Bottom left panel). Furthermore, the extended and mobile β- hairpin loop of SRS57 curved towards the center and guarded access to a potential groove (Figure 4.3C- Bottom right panel). An electrostatic surface representation of SRS57 likewise indicated that it lacks the basic surface groove that, in

SRS29B, is predicted to coordinate SPGs (Figure 4.3D).

4.2.4 Identification of T. gondii heparin-binding proteins

We next sought to confirm that native SRS57 from the parasite surface does not bind SPGs and to identify all Toxoplasma proteins capable of binding SPGs with strong affinity. Parasite lysates of RH and the Dsrs57 strains were subjected to heparin sulfate- agarose affinity chromatography. Fractions collected during the purification process were separated by SDS-PAGE and stained with Coomassie blue (Figure 4.4A). At least 10 bands were highly resolved by Coomassie staining of the eluate fraction (lane 4). To formally test whether any eluted proteins belonged to surface coat proteins SRS29B,

SRS34A or SRS57, the eluate fractions were subjected to immunoblot analysis using anti-

SRS29B (DG52), anti-SRS34A (5A6) and anti-SRS57 (1F12) monoclonal antibodies.

None of these proteins were recognized in the eluate fraction under the conditions utilized

(Figure 4.4B), indicating that these SRS proteins do not bind sulfated heparin with strong affinity. Probing the same fractions with chronic immune serum from mice infected with the Type II Me49 strain established that the 10 bands identified by Coomassie staining were immunogenic, parasite-derived, and possessed a strong affinity for sulfated heparin

(Figure 4.4C). Further studies will be necessary to identify these proteins and if they contribute to Factor H recruitment.

150 4.2.5 RHΔsrs57 parasites are not deficient in attachment, invasion, or extracellular survival kinetics

Because the crystal structure did not reveal an interaction between SRS57 and

SPGs, the functional relevance of SRS57 remained elusive. To further characterize these parasites, we wanted to determine if the lack of SRS57 impacted parasite survival, attachment, or invasion as previous work has suggested that SRS57 was necessary for infection competency (464). First, to assess extracellular survival, a time course experiment was performed. Wild type, ∆srs57 and complemented strains were subjected to extracellular conditions during 0, 0.5, 1, 2, 4, 8, 24 and 48 hours prior to infecting a confluent monolayer of human foreskin fibroblasts (HFF) cells with a dose of 103 tachyzoites/strain/time point. Analysis of the linear regression slopes showed no significant differences between the three strains, indicating that deletion of SRS57 had no impact on extracellular persistence (Figure 4.5A). Comparable results were obtained with the French

RH∆srs57 mutant CL55 and its complemented control in the assay conditions used, in contrast to published results for this knockout (Supplemental Figure 4.2). Similarly, a paired comparison between RH versus French RH strains (either wild type, knockout or complemented) showed no significant differences, regardless of the strain investigated

(Supplemental Figures 4.3A-4.3C). Time points of 24 and 48 hours were not presented because no plaque formation was observed, indicating that the parasites had a finite time in which they survived outside of the host cell.

Previous work had suggested that ∆srs57 mutants were defective in their ability to enter HFF cells (464). Plating efficiency was thus determined using a time course plaque

3 assay by inoculating HFF cells with 10 tachyzoites of each strain (RH, Dsrs57, and

151 complemented control). After 30, 60, and 90 minutes the monolayers were washed to remove uninvaded parasites. To determine the total number of invasion events possible in a 24-hour period (to serve as denominator for each individual strain tested) 250 tachyzoites were allowed to enter HFF cells. After 6 days, the number of plaques were counted and the percentage of plaques that formed compared to the 24-hour time point was calculated. No significant difference was observed between all strains tested (Figure 4.5B). Our findings were not in accordance with previously published work that used the French RH strain

(referred to as fRH, a Type I strain that is genotypically different from the RH strain isolated in the USA by Sabin in 1939). To determine if this was a peculiarity of the mutant clone generated using the fRH strain, we assayed the ∆srs57 CL55 mutant and its fRH parent. No difference was detected between fRH, the CL55 ∆srs57 clone or RH and the

∆srs57 clone 20 for 30 (Figure 4.5D) minutes, in the time course plaque assay performed in Figure 5B. When the complemented CL55 clone expressing wild type levels of

TgSRS57 was tested against RH and the ∆srs57 clone 20, no defect in extracellular survival was detected in vitro. (Supplemental Figure 4.3B) To assess if there was any defect in the rate of intracellular growth in HFF monolayers, the number of parasites per vacuole were counted after 16, 24- and 36-hours post-infection (Figure 4.5C). As determined for extracellular survival, the rate of growth did not differ significantly in the ∆srs57 parasites compared with the parental wild type and complemented strains.

4.2.6 In vivo, RHΔsrs57 infected mice show earlier cachexia and increased parasite burden, but no significant difference in survival

To determine if SRS57 contributed to pathogenesis in vivo, mice were first infected with Type I Δsrs57 parasites. Three groups of CD-1 mice were intraperitoneally challenged

152 (50 tachyzoites/mouse) with RH, Δsrs57 and complemented strains respectively. Survival and weight changes were monitored daily. Although there was no significant difference in survival among these groups, an unexpected early presentation of cachexia driven weight loss was detected by day 7 post infection in mice infected with Δsrs57 parasites compared with the other two groups, in which cachexia driven weight loss started at day 8 (Figure

4.6A). This was in contrast to previous studies in which the ∆srs57 strain CL55 failed to expand in vivo or cause weight loss. Strains used in those studies, fRH (French RH) and the mutant CL55 strain, were obtained, cultured and tested in mice using the same conditions used for RH and the ∆srs57 used herein. In agreement with this observation, bioluminescence imaging during the course of infection revealed a higher parasite burden beginning at day 6 post- infection in mice infected with Dsrs57 parasites (Figure 4.6B).

Quantification of the bioluminescence activity confirmed that the increase in parasite load was statistically significant (Figure. 4.6C). Intriguingly, these differences resolved during the following days and no significant difference in survival was observed (Figure 4.6D).

4.2.7 Type II Δsrs57 parasites are more virulent and exhibit higher parasite burdens in vivo

We next tested whether the absence of SRS57 in the CZ1Δsrs57 strain altered the virulence and parasite expansion kinetics in the Type II genetic background. C57BL6 female mice were infected with 2x103 parasites intraperitoneally. Survival and weights were monitored daily. Mice infected with CZ1Δsrs57 parasites died significantly (p=0.014) sooner than those infected with WT parasites (Figure 4.7A) and exhibited a more pronounced and earlier cachectic response (Figure 4.7B). These data confirmed the earlier

153 cachexia observed in mice infected with Type I RHΔsrs57 parasites and demonstrated that

SRS57 expression was protective in vivo.

To determine if parasite burden at day 6 post infection phenocopied the higher parasite burdens observed in mice infected with the RHΔsrs57 strain, peritoneal fluid and homogenized spleen and lungs were plated onto confluent HFF cells in 12 well plates and plaques were counted as a measure of parasite burden. Mice infected with Type II Δsrs57 parasites likewise demonstrated significantly higher numbers of parasites in all tissues assayed, demonstrating greater dissemination and parasite burden compared to mice infected with WT parasites (Figure 4.7C). These data suggest that SRS57 regulates pathogenesis by controlling parasite expansion.

4.2.8 Protection in vivo is dependent on complement but is not mediated by serum killing of parasites

C3 is a critical component of the innate immune system that limits infection by promoting direct lysis of pathogens or targets them for clearance by phagocytosis. To determine if the SRS57-mediated protection was C3-dependent, C3 deficient mice and WT mice were infected with WT parasites. Mice deficient in C3 were more susceptible to CZ1 infection than WT mice, demonstrating that complement was protective in vivo (Figure

4.8A). When the same mice were infected with the ∆srs57 parasites, which fail to bind WT levels of C3b, no protective effect was observed in mice replete with C3, suggesting that the interaction between SRS57 and C3 is mediating protection in vivo (Figure 4.8B). In addition, mice infected with Δsrs57 parasites died earlier than mice infected with WT parasites, and this was not dependent on C3 expression (Figures 4.8A, 4.8B). Hence, Type

154 II strains that are deficient in either SRS29C or SRS57, the two parasite proteins that are opsonized by C3, are more virulent.

Complement-mediated lysis is one mechanism of protection that can directly control pathogen proliferation. To test whether parasite resistance to serum killing was dependent on SRS57 expression, Δsrs57 parasites were incubated in 40% NHS for 30 minutes, and parasite viability and membrane attack complex formation (C5b-9) were measured by flow cytometry. Δsrs57 and WT parasites were equally resistant to human serum killing (Figure 4.8C) and had similar levels of C5b-9 formation (Figure 4.8D). In addition, mouse serum was previously shown to exhibit low complement activity and did not kill susceptible parasites as efficiently as human serum (Chapter 2). These data do not support complement mediated lysis as the major complement effector function controlling serum resistance, T. gondii infection load, or acute virulence in the mouse model. Hence, the opsonization of SRS29C or SRS57 appears to be a critical step in the host protective phenotype in vivo, These data suggest that other soluble factors associated with the complement pathway may be critical in promoting this protective phenotype.

4.2.9 SRS57 regulates systemic and local pro-inflammatory cytokine responses

Several studies have suggested that Type I parasites exhibit greater dissemination and parasite burdens compared to Type II strains in vivo, however dysregulated Th1 inflammatory cytokine production may also significantly contribute to mortality (342). To investigate if inflammatory cytokine production was a factor contributing to the altered virulence kinetics, C57BL/6 mice infected with 2x103 Type II Δsrs57 parasites intraperitoneally were sacrificed 6 days post infection. Serum and splenocytes were harvested and used to measure IL-12, IL-6, TNF-α, IFN-γ cytokine production. All

155 cytokines in the serum (Figure 4.9A) and those produced locally by ex vivo splenocyte cultures (Figure 4.9B) exhibited significantly higher levels in the absence of SRS57. These dysregulated cytokine responses could be explained by elevated parasite burden in the peritoneum, spleen and lungs compared to wild type mice (Figure 4.7C). Additional studies at earlier time points where parasite burdens are equivalent are required to determine if increased virulence of Δsrs57 parasites is driven by absolute parasite numbers or a failure to control parasite proliferation, or the result of a differential induction of the immune response in the absence of SRS57.

4.3 Discussion

Several host factors, including components of humoral immunity such as IgM

(466), complement proteins (173), and host sulfated proteoglycans (258), have been shown to interact with the tachyzoite surface coat. SRS proteins have been implicated in these interactions, however their contributions to T. gondii pathogenesis are not well understood.

In this study, we used reverse genetics strategy and a structural approach to investigate the specific interaction of surface protein SRS57 with SPGs, C3b and FH in order to gain a better understanding of its biological function. Here, we establish that SRS57 is a target for complement effector protein C3b. In contrast to SRS29C, the reduced capacity of SRS57 to interact with FH, however, could not be explained by interactions with SPGs. We established that SRS57 does not possess a lectin-specific activity for cell surface associated glycosaminoglycans, as previously suggested, and provided a structural blueprint to support our conclusion. Our data do confirm that at least 10 immunogenic parasite proteins specifically react with SPGs. Thus, it is possible that SRS57 interacts with other parasite proteins that bind SPGs and coordinate or present FH in a way that facilitates C3b

156 inactivation. Notably, in vivo characterization of the Δsrs57 parasites established that like

SRS29C, the interaction between SRS57 and C3 plays a critical role in regulating parasite virulence.

Microbial intracellular pathogens have evolved multiple strategies to gain entry to their target host cells. Whether the invasion process is passive or active, the initial recognition and attachment phase is critical for the success of internalization. A variety of cell surface effector proteins from both invading organisms and their hosts participate in this process. Many viruses, bacteria, fungi and parasites express lectins that bind to cell surface associated SPGs such as heparan sulfate to promote initial recognition, attachment and cell entry (462). It is also known that microbial pathogens utilize these interactions to evade and subvert host defense mechanisms, to their own advantage, in order to survive within the host (461, 467). SPGs are expressed in virtually all cell types, from simple invertebrates to humans (468). They are glycoproteins that contain one or more covalently attached linear polysaccharide chains referred to as glycosaminoglycans (GAGs). The two major types of GAGs found in animal cells are heparan and chondroitin sulfate (462).

Chondroitin sulfate A (CSA) is a cell surface receptor for Plasmodium falciparum-infected erythrocytes (469) and cytoadhesion of parasitized erythrocytes to the placenta is mediated by the specific interaction of a parasite encoded adhesin VAR2CSA with CSA present in placental proteoglycans (470). Plasmodium berghei uses the sulfation level of heparan sulfate proteoglycans (SPG) to navigate within the mammalian host. High levels of these glycoproteins on the surface of hepatocytes activate invasion, whereas low levels promote migration from the skin to the liver (471). The transition from a migratory to an invasive phenotype is mediated by a signaling pathway, which includes a calcium dependent protein

157 kinase 6. Binding of the sporozoite’s major surface protein, the circumsporozite protein

(CSP) triggers this process (471).

The first evidence that T. gondii possessed a sugar binding activity came from an early investigation in which it was shown that tachyzoites bind strongly to the neoglycoprotein bovine serum albumin-glucosamide (472). It was later shown that attachment of the parasite to its target cell is mediated by parasite lectins that bind to GAGs

(256). At least three surface adhesins that resolved at 26, 45 and 65 kDa by SDS-PAGE were identified for their lectin-specific activity and by their ability to bind to the surface of erythrocytes and HFF cells, although the molecular and biochemical characterization of these lectins, and their putative target sugars remain undefined (256). Despite the fact that the active process of internalization is conserved throughout the phylum, notable differences in the spectrum of invaded host cells is observed between different members of the phylum. Plasmodium spp. infect host hepatocytes and erythrocytes, whereas

Cryptosporidium spp. infect only enterocytes, and Toxoplasma possesses the unique characteristic of infecting virtually any nucleated cell (424). It is tempting to speculate that a correlation between host range and the number and variability of surface adhesins exists.

For example, more than 180 surface- associated SRS adhesin-encoding genes have been identified in the genome of Toxoplasma (261), whereas Plasmodium only possesses 14 members of the related 6-Cys protein family that, like Toxoplasma SRS proteins, are expressed developmentally, with different sets of 6-Cys proteins expressed at different stages throughout the parasite life cycle. Both families, the 6-Cys and SRS proteins, are derived from a common ancestor within the Apicomplexa and each have evolved their own adhesion superfamily of surface coat antigens based on distinct structural domains referred

158 to as the SRS and s48/45 domains in Toxoplasma and Plasmodium, respectively (272).

While relatively little is known about the specificity of the SRS proteins, three members of the analogous 6-Cys family play a role in gamete fertility (p230, p230p and p47) (473) although it is not clear whether these fertility factors interact via lectin binding. Other 6-

Cys proteins are known to facilitate invasion of hepatocytes (P52/P36) and promote binding to CD36 (Sequestrin) and Factor H (Pf92) (270–272, 474).

In addition to attachment and adhesion, evasion of host immunity is critical to establishing a successful infection. Studies in Chapter 2 demonstrated that SRS29C and

SRS25 contribute to serum resistance by recruiting host regulators Factor H and C4b- binding protein to downregulate alternative and lectin pathways of the complement system, respectively. The majority of serum resistance, however, was attributed to the regulation of the alternative pathway by Factor H. Factor H is composed of 20 homologous complement control proteins (CCP) domains, otherwise known as short consensus repeats, that give the appearance of flexible “beads on a string” that can fold back on itself (110).

Factor H predominantly interacts with C3b and host molecules such as sialic acids and

SPGs to discriminate between self and non-self surfaces. With several binding sites for proteoglycans and C3b, structural studies have established that Factor H simultaneously bind negatively charges anions like SPGs and the C3d portion of the C3b molecule (130) on host surfaces to mediate protection against complement. SRS29C fulfills both ligand requirements for AP regulation by its ability to bind both C3b and FH, however, our previous work showed that it was not the only target for C3b opsonization. Because

SRS29C exhibits strain-specific expression, we hypothesized that there were additional, more universal targets for C3b deposition on T. gondii strains. In this study, using reverse

159 genetics and a previously developed flow cytometric assay for measuring complement deposition, we identified SRS57 as another, and presumably the only other, C3b target.

Like SRS29C, significant loss of C3b deposition in the absence of SRS57 in both Type I and Type II strains indicated that it was a nucleophilic target for C3b. Analogous to

SRS29C, SRS57 also bound FH, exhibiting a partial reduction in its recruitment in its absence on Type II strains, indicating it may employ a similar SPG-dependent mechanism for recruitment. However, unlike S29C, it is unknown how SRS57 is recognized by complement recognition molecules, such as the mannose binding lectin, to facilitate C3b deposition. SRS57 lacks predicted N-glycosylation sites (toxodb.org, NetNGlyc1.0), thus it may not play a role in lectin pathway activation.

Previous studies have suggested that an array of T. gondii surface and secreted proteins, including SRS57, SRS29B, ROP2, ROP4, and GRA2 bind heparin (258, 264), a pharmaceutical mimic of heparan sulfate (HS) that is a highly-sulfated glycosaminoglycan, found in vivo in mast cells, and within connective tissue. It differs from HS in the degree of its sulfation, with heparin displaying higher N and O-sulfation than HS (475) and it is often used to study interactions between pathogens and host cells. A homology model of

SRS57 based on the structure of SRS29B suggested a possible SPG- binding ability for

SRS57 through a positively charged binding cleft (265). In order to verify this model and to investigate the structural and molecular basis of SRS57-SPG interaction, we determined the high-resolution crystal structure of SRS57. We found that SRS57 shared the same architecture features with previously characterized SRS proteins. However, in contrast to the homology model, our structural data did not identify a positively charged binding cleft capable for accommodating SPG. Moreover, our binding studies with native or

160 recombinantly expressed SRS57 failed to indicate any binding with heparin. Notably, binding studies in the past showing SRS57-SPG interactions (258) had made use of recombinantly expressed His-tagged proteins and the presence of the 6×His may non- specifically bind to heparan sulfate (HS) (476), giving rise to false positive results.

In the absence of identifying SPGs bound to SRS57, we sought to identify the SPG binding proteins using heparin sulphate- agarose affinity chromatography to identify other potential surface coat proteins that might facilitate FH recruitment and retention to the parasite surface. In total, at least 10 high affinity SPG binding proteins were identified.

Consistent with our structural and functional analyses, we purified native SRS57 from the parasite surface and failed to identify any interaction with heparin. Taken together, these results implied that SRS57 recruitment was not SPG-dependent, indicating there must be an alternative mechanism to sequester FH at the parasite surface in an SRS57-dependent manner. Interestingly, several microbes are known to bind FH domains 19-20, which are the same domains that bind to heparin, and heparin is known to inhibit FH binding to these microbes. (477). However, the molecular basis for this common microbial binding site is poorly understood. Further characterization of the FH domains that specifically bind to the parasite surface would determine if T. gondii similarly uses this common binding site.

Alternatively, it cannot be ruled out that the C3b-modified SRS57 forms a complex with, or is in close proximity to, one or more of the 10 immunogenic SPG-binding proteins identified in this study. Members of the SRS family possess at least four conserved cysteine residues that are known to disulfide bond and mediate the appropriate folding and dimerization of the nascent protein (265). It is possible that it heterodimerizes with another

SRS protein that has lectin activity, as the structurally related 6-CYS proteins from

161 Plasmodium are known to heterodimerize (474), but thus far, no evidence exists for SRS heterodimerization. The flexible structure of the Factor H molecule (110) which is retained upon binding C3b (478), and its multiple SPG (121, 128, 479, 480) and C3b (481, 482) recognition domains support the possibility of multivalent interactions (483). This flexibly would allow for several parasite factors to facilitate the formation of a tripartite complex with C3b and SPG to promote the recruitment of FH, however additional studies will be required to identify the precise molecular details that promote FH binding to SRS57.

Because the functional relevance of SRS57 remained elusive, we further characterized SRS57 deficient strains in vitro and in vivo. The SRS57 deficient strain that we generated in the type I RH background, and the previously published CL55 ∆srs57 mutant, both failed to identify any difference in extracellular survival, adhesion, intracellular replication or a change in murine virulence. This is in stark contrast to the previously published results for this knockout which indicated a role for SRS57 as an adhesin facilitating the invasion process and virulence in mice (258, 464). Although we did observe a difference in dissemination kinetics in vivo for the SRS57 deficient Type I strains, this did not result in a dramatic change in the kinetics or time to death. While it is formally possible that the differences observed in host susceptibility between the two studies are the result of the type of mouse strain used, as it has been shown previously that host genetics can influence both acute mortality and cyst load (484), the fact that our infections used the more resistant CD-1 outbred strain whereas the previous work was done using susceptible inbred Balb/C lines argues against this as a likely explanation.

We also generated a Dsrs57 strain in a less virulent Type II CZ1 strain and infected susceptible C57BL6 mice to further investigate the differences in parasite burden and

162 cachexia observed in the Type I deficient strain. The CZ1Dsrs57 parasites were significantly more virulent which was opposite to the protective phenotype previously observed for Dsrs57 parasites (464). However, this phenotype was similar to the observed for the CZ1∆srs29C mutant, the other SRS protein known to be modified by C3b.

Furthermore, these mice exhibited higher parasite burdens and, like the CZ1∆srs29C mutant, promoted a more vigorous Th1 pro-inflammatory response, suggesting that the covalent modification of SRS57 with C3b attenuates virulence in a manner similar to

SRS29C by regulating parasite proliferation during acute infection. However, it is not clear from these studies if uncontrolled parasite proliferation or immunopathology contributed to the rapid mortality of mice infected with Dsrs57 parasites.

Because SRS57 was a target for C3b, the biological relevance of C3 during acute infection was evaluated next. C3-/- mice infected with SRS57 sufficient WT parasites were more susceptible to infection, indicating that complement was also protective during acute infection. Although no difference in virulence between wild type mice and C3-/- with

Dsrs57 parasites was observed, C3-/- mice infected with Dsrs57 parasites succumbed to infection much sooner (day 12) than C3-/- mice infected with wild type parasites (day 20), suggesting there is a factor independent of C3 that may be interacting with SRS57 to synergistically regulate virulence in vivo. The complement system is known to cross talk with several other systems, including TLR receptors signaling (485, 486) and C-type lectin receptor Dectin-1 (487), to coordinate immunological responses (488), thus it is possible

SRS57 might coactivate complement and one of these systems to regulate virulence and control parasite proliferation.

163 Since SRS57 expression did not impact complemented-mediated lysis and we previously demonstrated that mouse complement does not promote parasite killing of serum-susceptible parasites either in vitro or in vivo (Chapter 2), it is likely that additional complement effector functions, such as complement-mediated phagocytosis or stimulation of adaptive immunity, was affected by the reduction of iC3b deposition on Dsrs57 parasites or in the case of C3 deficiency. Interestingly, loss of complement mediated protection in these mice occurred at the induction of the adaptive immune response (before day 21), indicating that complement might be mediating its effects by regulating adaptive immune responses. Both C3b and anaphylatoxins C3a and C5a play a role in regulating B and T cell responses, respectively. C3a and C5a have recently been shown to play a critical role in regulating Th1 responses (152, 158), which is an essential response for resolving acute

T. gondii infection. Western blot analysis showed that the reduction in FH recruitment impacted the amount of iC3b on Dsrs57 parasites, which can also alter the amounts of C3a and C5a being produced that can subsequently modulate cytokine production and possibly

T cell responses. Alternatively, antigen covalently bound by catabolite C3b, C3dg, are recognized by CR2 on B cells and lower the threshold for B cell activation and antibody production (143, 144). In the absence of SRS57 or complement, it is possible that these mice exhibit altered SRS57-specific B cell responses that result in less effective B cell immunity. Further studies are required to determine which C3 effector proteins contribute to the enhanced mortality in infected C3 deficient animals and what role SRS57 and

SRS29C plays in such immune modulations.

In aggregate, our data indicate that unlike SRS29C, SRS57 does not possess a lectin-specific activity nor does it mediate attachment through an SPG host cell receptor

164 interaction, indicating FH recruitment by SRS57 is SPG-independent. However, at least 10

T. gondii lectins were identified that specifically interact with heparin. Ongoing studies to identify the nature of these proteins will shed light into the mechanisms of initial host cell recognition and the possible mechanism for SRS57-dependent FH recruitment. In vitro and in vivo studies revealed a novel function for SRS57 via its interaction with C3b, which contributes to its biological function by regulating parasite virulence and promoting parasite persistence in the mouse model.

4.4 Materials and Methods

4.4.1 Parasite strains, host cells and cell culture conditions

Wild type RH and CZ1 tachyzoites, knockout and complement strains were propagated in human foreskin fibroblasts (HFF) in Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% heat inactivated fetal bovine serum (FBS), 2 mM glutamine, 10

μg/ml of gentamicin and 0.5% of penicillin/streptomycin (100X) and maintained at 37oC in a humid 5% CO2 atmosphere.

4.4.2 Targeted disruption and complementation of the SRS57 locus

The targeting vector used to generate the Δsrs57 strain was derived from the pminiHXGPRT vector (Catalogue Number: 2855. NIH AIDS Reagent program,

Contributor: David Roos). The 5’ flanking (flk) sequence (2.0 Kb) and 3’ flk sequence (3.1

Kb) were amplified from RH genomic DNA and cloned into KpnI/HindII and BamH1/SacI sites, respectively. The primers to amplify the 5-flanking sequence were 5SRS57FW

(GCGGTACCAAGTCGAAGA GTGCGTTCGT) and 5SRS57Rev (GCAAGC

TTAAGGATACCGTGTGCGAAAAC) and the primers to amplify the 3’-flanking sequence were 3SRS57FW (GCGGTACCGT GTGAGACCCTCTCT) and 3SRS57Rev

165 (GCAAGCTTACGAATCCAGTGCAGTTTCC). The 5’dhfr-HXGPRT-3’dhfr cassette

(1.9 Kb) contained in the vector was used as a marker for positive selection. The 5’flk

TgSRS57/dhfr-HXGPRT-dhfr/3’flk SRS57 construct (7.0 Kb) was released from the vector using the restriction enzymes KpnI and SacI. 107 RHΔhxgprt parasites tachyzoites were transfected with 50 μg of the construct. Transfections were performed by resuspending parasites and DNA in electroporation buffer (120 mM KCl, 0.15 mM CaCl2, 10 mM

K2HPO4/KH2PO4. pH 7.6, 25 mM HEPES, pH 7.6, 2 mM EDTA, 5 mM MgCl2, 2 mM

ATP, 5 mM glutathione), using a BTX model ECM630. 24 hrs after transfection parasites were selected with 50 μg/ml mycophenolic acid and 50 μg/ml xanthine. Surviving populations were cloned by limiting dilution and screened for the loss of the TgSRS57 expression by flow cytometry. For complementation, 107 parasites knockout were co- transfected with 50 μg of the TOPO 4.0 vector harboring a PCR amplified SRS57 genomic locus including the promoter and the 5’ and 3’ untranslated regions (3.7 Kb) plus 5 μg of a pGFP-LUC plasmid, containing cDNA sequences of the Green Fluorescence Protein

(GFP) and Fire Fly Luciferase (LUC). The primers used for the amplification of the SRS57 locus were PF: CAAGACACACAAGTGCGTGTGATG AC and PR:

CACTGTTCTGATGGAATGCT CGCTAC and TA cloned into the TOPO 4.0 vector.

Before transfection both plasmids were linearized with Not1. 10 days post- transfection, stable parasites expressing GFP were FACS sorted onto 96 well plates. Positive clones expressing GFP were analyzed for TgSRS57 surface expression by flow cytometry.

166 4.4.3 Generation of CZ1Δsrs57 mutant strain using CRISPR/Cas system

A clonal isolate of CZ1-Δku80/Δhpt was used to generate the CZ1-Δku80/Δsrs57 parasites. The Srs57 gens was replaced by the drug selectable marker HPT (HXGPRT – hypoxanthine-xanthine-guanine phosphoribosyl transferase) flanked by 40bp homologies to SRS57 using CRISPR/Cas system (SRS57 guide RNA sequence:

GTGTGTGTTGTCTGCGATCT). Fifteen micrograms of HXGPRT repair oligo and 30 μg of guide RNA were transfected into by electroporation using the BTX system. The parasites were then allowed to infect T25 flask of HFFs for overnight, after which the medium was changed to mycophenolic acid (MPA) and xanthine at 50ug/ml for HXGPRT positive selection. After two passages, the parasites were cloned using single-cell method into 96 well plates. Δsrs57 clones were identified by PCR of the integrated of HXGPRT into the open reading frame using the following primers: (SRS57 forward primer:

CTTTTTCAGTAACGCTTCGT and HXGPRT reverse primer:

GAGCACCCTGATATGACAAT). SRS57 deletion was further confirmed by staining and analyzed by flow cytometry and by western blot.

4.4.4 Flow cytometry

Suspensions of freshly harvested tachyzoites from the different strains were washed with

PBS, fixed for 15 min with 2% (w/v) formaldehyde in PBS at room temperature (RT) and stained with the primary monoclonal antibody 1F12 (specific for the SRS57 protein), diluted 1:4000, washed three times with FACS buffer (1% BSA in PBS) and stained with the secondary antibody anti-mouse conjugated with Allophycocyanin (APC), diluted

1:200. After 3 washes in FACS buffer, the stained parasites were analyzed for surface

167 protein expression by flow cytometry. Data were collected using a FACS Canto flow cytometer (BD Bioscience) and analyzed using FlowJo software.

4.4.5 Flow cytometry-based complement assays for C3b deposition, FH recruitment,

C5b-9 formation and parasite viability

Complement deposition was measured by flow cytometry as previously described (Chapter

2). Briefly, parasites were collected from a freshly lysed monolayer of human foreskin fibroblast cells, filtered, and washed twice with PBS to remove media. 1x106 parasites per time point were incubated with 10% non-immune human serum (pooled NHS, Cederlane)

++ in complement activating buffer HBSS (Hanks Buffered Saline Solution, 1mM MgCl2,

0.15mM CaCl2). Complement activation was stopped using cold PBS and washed twice with cold PBS to remove excess serum. For viability assays, parasites were exposed to 40%

NHS, washed three times with cold PBS to remove excess serum, and stained with a fixable viability stain (Life Technologies) for 15 minutes and washed twice with PBS. Parasites were then fixed with 1% PFA solution for 10 minutes, washed twice, and stained with mouse α-human C3b/iC3b antibody (1:500) (Cedarlane), mouse α-human C5b-9 (1:500),

(Santa Cruz Biotechnology) or goat α -human Factor H (1:2,000) (CompTech), followed by α-mouse APC (eBiosciences) or α-goat APC conjugated antibody (Jackson

Laboratories), both at 1:1,000. Antibodies were diluted in FACs buffer (PBS + 1% BSA), incubated for 30 minutes at room temperature and protected from light, followed two washes with FACs buffer. 20,000 events were collected on a BD Fortessa flow cytometer and analyzed by FACs DIVA, FlowJo and Prism software.

168 4.4.6 Extracellular parasite survival

Suspensions of free parasites (10 tachyzoites/ml) were incubated in culture media at 37°C with 5.5% CO2, in absence of host cells for 0, 1/2, 1, 2, 4, 8, 24 and 48 hrs prior infection.

After the indicated times a total of 103 tachyzoites of each strain was applied to a fresh monolayer of HFF cells, seeded onto 24 well plates and incubated during 5 days in normal culture condition without disturbance. After incubation, the wells were rinsed with PBS, fixed with 100% of methanol for 5 min and stained with crystal violet. Parasites survival was estimated by counting the number of plaques. The assay was carried out in triplicates.

The results were presented as the mean ± SD (n=3).

4.4.7 Plating efficiency

103 freshly lysed tachyzoites were allowed to invade monolayers of HFF cells for 30, 60, and 90 min and 24hrs. After the indicated times, uninvaded parasites were removed by washing four times with PBS and allowed to replicate undisturbed at 37oC and 5.5% CO2 atmosphere for 6 days. The total number of plaques formed at each time point were counted. The plating efficiency was expressed as a percentage of the total number of plaques at 24 hrs. The experiment was performed in triplicate and the statistical differences among means were calculated using the One-way ANOVA (non-parametric) test.

4.4.8 Rate of replication

Confluent monolayers of HFF cells, grown in 12 well plates, were infected with 105 tachyzoites/well. After 4 hrs of incubation at 37oC and 5.5% CO2, to allow the parasites to invade, the wells were washed 3 times with PBS and incubated for 16, 24, and 36 hrs in regular media. At each different time the cells were fixed with 100% of methanol for 10 min and the average number of parasites/vacuole was scored by counting 100 randomly

169 selected vacuoles. The results are presented as the mean of two independent experiments

± SD.

4.4.9 Mouse infections

Outbred female CD-1 mice (from Charles River), female C56BL/6 mice (Jackson), and female C3-/- mice (Jackson), 6-8 weeks old, were used for in vivo experiments. For infections with RH strains, groups of 5 mice of each strain were intraperitoneally infected with 50 freshly harvested tachyzoites. For CZ1 strains, 2 x103 freshly harvested parasites were injected intraperitoneally into wild type C56BL/6 mice and C3-/- mice. The cachexia induced during the course of the acute infection was monitored daily. All procedures were performed in accordance with the guidelines of the National Institutes of Health.

4.4.10 In vivo imaging

For living image, mice were injected with 150 μl (3 mg) of D-luciferin substrate, maintained at least 10 min to allow dissemination of the substrate and anesthetized in an oxygen-rich induction chamber with 2% isofluorane. Images were collected during 5 min using an IVIS (In Vivo Imaging System) Spectrum (PerkinElmer). Mice were imaged at days 5, 6, 7, and 8 post infection. Data acquisition and analysis was performed using the living imaging software.

4.4.11 Plaque assays to quantify parasite load

Mice infected with CZ1 strains were euthanized at day 6 post infection. Peritoneal fluid was collected by injecting 5ml of sterile PBS into the peritoneal cavity of euthanized mice.

Spleens and lungs were harvested and homogenized over 70-μm strainers in 5ml of X-

VIVO-20 serum free media. Ten-fold serial dilutions starting with 500 μl of resuspended homogenate were added to HFF monolayers in 12 well plates and cultured undisturbed in

170 DMEM media for 7-9 days. HFFs were fixed in methanol and stained with crystal violet.

Plaques were counted using an inverted microscope and parasite loads were expressed as

CFU per organ.

4.4.12 Ex vivo splenocyte cultures

Spleens were homogenized over 70-μm strainers in 5ml of X-VIVO-20 serum free media.

After red blood cell lysis (Lonza ACK lysis buffer) splenocytes were plated in a 96-well flat bottom plate at 5x105 splenocytes per well in 250μl of X-VIVO serum free media

(Lonza) and cultured for 24 hours. 250μl of supernatants were collected for analysis.

4.4.13 Measuring cytokine production from serum and ex vivo splenocyte by ELISA

After euthanasia, sera were prepared from blood collected by cardiac puncture. Blood was allowed to clot for 30 minutes and then spun at 1000rpm for 15 minutes to extract serum.

Supernatants of ex vivo splenocyte cultures were collected as described above. Samples were analyzed using ELISA kits for IL-12p40, IFN-γ, TNF-α, IL-6, IL-18 and IL-10 according to manufacturer’s instructions (R&D Systems).

4.4.14 Cloning, expression and purification of SRS57

The SRS57 sequence from the signal peptide cleavage site (SP) to GPI anchor site (GPI) was codon optimized, synthesized by GenScript, and subcloned into a modified pAcGP67b vector (Pharmingen) incorporating a C-terminal hexahistidine tag and thrombin cleavage site. SRS57 construct extending from the end of the predicted signal peptide to the end of predicted D2 was cloned using PCR. SRS57 clones were transfected with linearized

Baculovirus DNA into Sf9 cells and amplified to a high titre. Hi-5 cells at 1.8 x 106 cells ml-1 were infected with amplified virus for 65 hrs, after which time the supernatant was harvested, concentrated and applied to a HisTrapFF Ni affinity column. SRS57 was eluted

171 with an increasing concentration of imidazole and fractions analyzed by SDS- PAGE and pooled based on purity. The hexahistidine tag was removed by thrombin cleavage and

SRS57 was further purified by size exclusion chromatography (Superdex 16/60 75) in

HEPES buffered saline (HBS – 20 mM HEPES pH 7.5, 150 mM NaCl).

4.4.15 Crystallization and data collection

Diffraction quality crystals of TgSRS57 were grown in 25% PEG1500. The final drops consisted of 0.6 μl protein (12 mg ml-1) with 0.6 μl reservoir solution and were equilibrated against 100 μl of reservoir solution. Cryo protection was carried out in mother liquor supplemented with 25% glycerol for 20 seconds and the crystal was flash cooled at 100 K directly in the cryo stream. Diffraction data were collected on beamline 9-2 at SSRL

(Stanford Synchrotron Radiation Laboratory).

4.4.16 Data processing, structure solution and refinement

Diffraction data to a resolution of 1.59 Å were processed using Imosflm (452) and Scala

(489) in the CCP4 suite of programs (490). Initial phases were obtained by molecular replacement (MR) using PHASER (491) with the individual D1 and D2 domains of

TgSRS29B (PDB 1KZQ) pruned with CHAINSAW (492). Solvent molecules were selected using COOT (46) and refinement carried out using Refmac5 (493). Stereo- chemical analysis performed with PROCHECK and SFCHECK in CCP4 (490) showed excellent stereochemistry with more than 97% of the residues in the favored conformations and no residues in disallowed orientations of the Ramachandran plot. Overall 5% of the reflections were set aside for calculation of Rfree. Data collection and refinement statistics are presented in Table 1. The atomic coordinate and structure factor files have been deposited in the PDB under accession code 5WA2. Electrostatic surface images were

172 calculated using Adaptive Poisson-Boltzmann Solver (APBS) electrostatic surface evaluation tool with default parameters (Ionic concentration of 0.150 mol/l NaCl, biomolecular dielectric constant of 2 and solvent dielectric constant of 78.54 (water). The charges and radii of atoms in the PQR file required by APBS were taken from AMBER94 force field (494).

4.4.17 Heparin-binding assay

To isolate heparin binding proteins, 108 freshly harvested tachyzoites were washed with

PBS and then lysed for 30 min on ice in 100 μl of lysis buffer (1% of NP-40, 0.25M NaCl in phosphate buffer pH 7.2). The lysates were centrifugated at 14.000 rpm for 30 min.

Cleared lysates were applied to a heparin- agarose type I beads (Sigma) previously equilibrated with 0.2% NP-40 in phosphate buffer, pH 7.2 and incubated for 30 minutes under gently rotation. Bound beads were extensively washed with 0.2% NP-40, 0.5M NaCl in phosphate buffer pH 7.2, followed by elution of the heparin bound proteins with a high salt elution buffer (0.2% NP-40, 1M NaCl in phosphate buffer pH 7.2).

4.4.18 SDS-PAGE electrophoresis and Western blot

Lysate, flow through, last wash and eluate fractions were analyzed by SDS-PAGE on

Novex bis-tris 4 to 12% gradient gels (Invitrogen) using MOPS SDS as running buffer.

Samples were heated at 70oC for 10 min in sample buffer containing 50mM dithiothreitol as reducing agent. Gels were stained with Coomassie Blue. For Western blot, the gels were electro transferred to nitrocellulose over night at 10V constant, using a Mini Trans-Blot®

Electrophoretic Transfer Cell (BioRad). After transfer, the membranes were blocked in

PBS containing 5% non-fat dry milk and 0.05% Tween 20 prior the addition of the appropriate antibodies (anti-SRS29B (DG52) mAb, 1:5000; anti-SRS34A (5A6) mAb,

173 1:400; anti-SRS57 (1F12) mAb, 1:1000, or mouse chronic serum after 5 weeks post infection. After three washings with PBS containing 1% non-fat dry milk and 0.05% Tween

20, the membranes were incubated with peroxidase conjugated goat anti-mouse antibody

(Sigma), diluted 1:1000), washed three times and visualized by chemiluminescence.

Alternatively, parasites extracts were treated with GPI-specific phospholipase C (GPI-

PLC). Briefly, parasites were washed with PBS, resuspended in 50 ml of lysis buffer (50 mM Tris [pH 8.0], 5 mM EDTA, 5% NP-40) and incubated with GPI- PLC from Bacillus cereus for 1 h at 37°C. Cross-reacting determinants (CRDs) of GPI anchors, which were unmasked by this treatment, were detected with a polyclonal antiserum against CRD.

For C3b detection, parasites were treated with 10% NHS for 20 minutes, washed three times with cold PBS, and lysed with 1% NP-40 lysis buffer (1% NP-40, 150mM

NaCl, 50mM Tris pH 8.0) for one hour on ice and pelleted at 14,000 rpm for 20 minutes to remove insoluble material. Lysates were prepared with 2X Laemmli sample buffer

(Biorad) with 5% 2-mercaptoethanol and boiled for 5 minutes at 95°C. Samples were loaded onto 12% polyacrylamide gels (Biorad) and transferred onto nitrocellulose using

GenScript eBlot system. Membranes were blocked for 1 hour with 5% non-fat dry milk in

PBS + 0.05% Tween-20 (PBS-T). The following antibody dilutions were used: rabbit α-

SAG1 1:1,000, goat α-C3b (CompTech) 1:20,0000 and incubated at room temperature for

1 hours. Membranes were washed three times with PBS-T and incubated with α- goat HRP

1:5,000 (Santa Cruz Biotechnology, Inc.) for one hour, followed by three washes with PBS-

T. Proteins were detected with Clarity Western ECL Substrate (Biorad).

174 4.5 Figures

Figure 4.1. Targeted deletion of the SRS57 gene. (A) (Top) Diagram of a genomic fragment spanning the SRS57 genomic locus. The SRS57 coding sequence is indicated by a black box. SP indicates the signal peptide and GPI the glycosylphosphatedylinositol anchor. 5’ and 3’ UTRs are indicated by white boxes. (Bottom) 5’dhfr-HXGPRT-3’dhfr construct used to disrupt the SRS57 gene by homologous recombination. (B) Anti-CRD Western blot of RHWT (lane 1), RH TgSRS57 mistargeted (lane 2) and 3 independent clones of RHDsrs57 strain (lanes 3, 4 and 5), showing the absence of TgSRS57. Tachyzoites pellets of each strain, were solubilized in 1% NP-40, treated with GPI-PLC, separated by SDS-PAGE, transferred to nitrocellulose and probed with anti-CRD polyclonal antibody. (C) Flow cytometry analysis of RHWT (black), RHDsrs57 (white) and complemented (dark grey) strains labelled with anti-SRS57 (1F12 monoclonal antibody) on the left and anti-TgSRS34A (5A6 monoclonal antibody) on the right.

175 Expression of SRS57 was completely abolished in the knockout strain, whereas the expression of TgSRS34A was not affected. (D) Schematic representation of knockout construct for SRS57. The endogenous locus was disrupted using the hypoxanthine- xanthine-guanine phosphoribosyl transferase (HXGPRT) selectable marker and CRISP methodology. (E) Flow cytometry analysis of CZ1 wild type (black) and Dsrs57 (gray) labeled with anti-SRS57 (1F12 monoclonal antibody). (F) Western blot analysis of 1x106 CZ1 wild type strain and Dsrs57 parasites. Nitrocellulose membranes were probed with anti-SRS57 (1F12 monoclonal antibody). The RHDsrs57 strain was generated by Dr. Viviana Pszenny, NIH and the CZ1Dsrs57 strain was generated by Gloria Adedoyin, NIH.

176 Figure 4.2. SRS57 expression impacts complement deposition. C3b deposition time course of (A) RH wild type (black square) and RH Dsrs57 (open square) and (B) CZ1 (black hexagon) and CZ1Dsrs57 (open hexagon) incubated in 10% non-immune human serum (NHS) over 60 minutes at 37°C. Serum was diluted in complement activation buffer, Hanks Buffered Saline Solution supplemented with 0.15mM CaCl2 and 1mM MgCl2 (HBSS++). Wild type parasites in 10% heat inactivated serum (Δ56) were used a negative control (open square, dotted line). C3 deposition was measured by staining parasites with a monoclonal mouse α-human C3b/iC3b antibody (1:500, Cedarlane). Flow cytometry analysis of Factor H recruitment on (C) Type I RH (black square) and RHDsrs57 (open square) or (D) Type II CZ1 (black hexagon) and CZ1Dsrs57 (open hexagon) parasites incubated in 10% NHS over 60 minutes at 37°C. Wild type parasites in 10% heat

177 inactivated serum (Δ56) were used a negative control (open square, dotted line). Factor H was measured by staining parasites with goat anti-human Factor H (1:2,000, Complement Technologies). (E) Western blot analysis of C3b deposition on RH and RHDsrs57 parasites incubated in 10% NHS for 20 minutes at 37°C. Membranes were probed with goat anti- human C3b at (1:20,000, Complement Technologies). Error bars represent mean ± SEM. Time course experiments and western blot are representative of at least two experiments.

178

Figure 4.3. Structural divergence of the SRS fold and biological implications of the SRS57 crystal structure. (A) Top: Schematic representation of SRS57 with predicted structural domains, GPI regions and signal peptides (SP). Orange rectangle represents the crystallization construct. Below- SEC profile of SRS57 (orange line). Inset, Coomassie- stained SDS-PAGE gel of SRS57 (expected molecular weight: 32 kDa). The protein standards used were conalbumin (75 kDa; peak I), ovalbumin (43 kDa; peak II), and carbonic anhydrase (29 kDa; peak III. (B) Left: Tertiary structure showing the head to tail structure of SRS57 in the predicted orientation with respect to the parasite cell surface with the N- terminal domain (D1) in bright orange and the C- terminal domain in light orange. The disulphides are indicated as yellow sticks. The C- terminal tail is indicated in light orange. Inset: 2Fo − Fc σA weighted electron density maps at 1.2σ showing the interdomain linker region (C). Top: Topology diagrams of the β strands that form upper leaves of D1 in SRS57 (orange), SRS16C (purple), and SRS29B (cyan). SRS57 is most similar to SRS16C and notably more substantial than SRS29B. The red arrows indicate the extra β strand found in SRS57 and SRS16C whereas the black arrow indicates the extended β

179 stands in SRS57. Bottom: Left- Superimposition of SRS29B dimer (PDB id: 1KZQ) and SRS57 showing the first β stand (red arrow) of TgSRS57 that clashes with the other monomer (dark teal) of SRS29B dimer, thereby preventing dimerization. Right- Side view of the structural overlay showing the curved β- hairpin loop (dotted box) that impedes access to the potential groove. Inset- B-factor putty model depicting the mobility of the hairpin loop. (D) Top: Top view of the SRS29B dimer highlighting the basic groove, Bottom: Same top view of SRS57 and SRS29B monomer, along the SRS29B dimer axis. Electrostatic surface is colored as calculated by APBS (48) (−1 kT/e in red and +1 kT/e in blue); k, Boltzmann’s constant; e, elementary charge. Crystal structure of SRS57 was solved by the group of Martin Boulanger, University of Victoria.

180

Figure 4.4. Detection of T. gondii heparin-binding proteins. Pellets of tachyzoites from wild type and knockout strains were lysed in lysis buffer containing 1% NP-40. Insoluble material was collected by centrifugation and the supernatant was subject to heparin-agarose affinity chromatography. The different fractions were analyzed by SDS-PAGE and Western blot. (A) Coomassie blue stained SDS-PAGE showing the protein profile of the fractions corresponding to the process of heparin-agarose affinity chromatography. Lane 1, loading sample. Lane 2, unbound flow through fraction. Lane 3, wash of the heparin column. Lane 4, high NaCl eluate from the affinity column. Both eluates (WT and knockout) show identical protein profile. (B) Immunoblot of the samples in (A) probed with anti-TgSRS29B (top), anti-TgSRS34A (middle) and anti-TgSRS57 (bottom). None of these three SRS proteins bound to heparin. (C) Immunoblot of the samples in (A) developed with chronic serum from mice infected with T. gondii. A wide range of proteins were observed in the eluate fraction, indicating the no binding observed in panel (B) was genuine. This assay was performed by Dr. Viviana Pszenny, NIH.

181

Figure 4.5. In vitro characterization of extracellular survival, attachment/invasion, plating efficiency and intracellular growth of Δsrs57 parasites. (A) Quantification of the extracellular survival. 103 tachyzoites of each strain (RH WT, Δsrs57 and complement) were maintained for 0 min, 30 min, 1, 2, 4 and 8hrs in extracellular condition, before applying to a fresh monolayer of HFF cells. After 5 days, undisturbed in normal culture conditions, the number of plaques formed at each time point were counted. No significant differences were observed in the extracellular survival and plating efficiency between the strains. Values represent the mean of triplicates ± SD. (B) Time course plating efficiency. 103 tachyzoites of each strain (RH WT and Δsrs57 [clones 20 and 27], and complemented Δsrs57) were inoculated onto monolayers of HFF cells, grown in 24 well plates, and allowed to invade for 30, 60, 90 min, and 24 hrs. After extensive washes, the parasites were incubated undisturbed in normal culture conditions and the total number of plaques were recorded after 6 days. The results were expressed as a percentage of total number of plaques counted at 24 hrs. Data represent the mean of 3 independent experiments ± SD. The statistical significance was analyzed by One-way ANOVA (non-parametric) test, using Prism 7 software. The results show no significant differences between strains, indicating that the lack of TgSRS57 does not affect the ability to invade. (C) Rate of replication. Confluent monolayers of HFF cells, were infected with 105 tz/well of each strain and incubated for 16, 24, and 36 hrs. The average number of parasites/vacuole was scored by

182 counting 100 randomly selected vacuoles. The results are presented as the mean of two independent experiments ± SD. No significant differences in in vitro intracellular proliferation were observed between RHWT, Δsrs57 and the complemented strains. (D) Comparison of plating efficiency between RH and fRH strains. No significant differences 3 in plating efficiency were observed between strains. 10 tachyzoites of each strain (RH WT, ∆srs57 [clone 20], fRH and fRH∆srs57 [clone CL55]) were inoculated onto monolayers of HFF cells grown in 12 well plates, allowed to invade for 60 min, washed extensively, and incubated undisturbed in normal culture conditions. The total number of plaques was recorded after 6 days. The results were expressed as a percentage of total number of plaques counted at 24 hrs. Data represent the mean of 3 independent experiments ± SD. The statistical significance was analyzed by One-way ANOVA (non-parametric) test using Prism 7 software. This assay was performed by Dr. Viviana Pszenny, NIH

183

Figure 4.6. In vivo characterization of Type I Δsrs57 parasites (A) Loss of weight after murine infection. For each strain, groups of 5, 4 or 3 CD1 mice were intraperitoneally infected with a dose of 50 parasites/mouse and the daily weight was registered until death. The results correspond to one of three independent experiments. Earlier cachexia was observed in Δsrs57 parasites, started at day 6 post infection, compared with RH WT and complement strains, started at day 7 post infection. The curves represent the mean ± SD of 5, 5 and 3 animals infected with RH WT, Δsrs57 and complement strains respectively. (B) Bioluminescent detection of parasite burden in vivo. CD-1 mice were infected with 50 luciferase-positive WT, Δsrs57 or complemented strains and imaged daily. Representative images of 3 animals per group, with photon output intensity in photons/second/cm2/surface radiance are shown for days 5, 6, 7, and 8. The image shows higher luciferase intensity at day 6 in mice infected with Δsrs57 parasites, compared with the WT and complemented strains. (C) Quantification of the luciferase activity showing that the differences in parasite burden observed at day 6 post infection, in the Δsrs57 parasites, was statistically significant (p<0.05) confirming the visual increases in luciferase activity as observed in (B) at day 6 and consistent with the loss of weight as shown in (A). (D) Survival curve of CD1 mice challenged with 50 tachyzoites/mouse of the indicated strains. No significant differences were observed between the 3 strains. The results are an aggregate of three independent experiments.

184

Figure 4.7. Mice infected with Type II Δsrs57 parasites exhibit altered pathogenesis. (A) Survival curve of C57BL/6 mice infected with 2x103 tachyzoites intraperitoneally. Mice infected with Δsrs57 parasites (teal line) died significantly sooner than those infected with wild type parasites (black line) (p = 0.014). The results are an aggregate of two independent experiments, n=9 per group. (B) Weight loss after infection with 2x103 tachyzoites intraperitoneally. Earlier cachexia was observed in Δsrs57 parasites, starting at day 3 post infection, n=9. (C) Parasite burden was determined at day 6 post infection by plaquing peritoneal fluid and homogenized spleens and lungs on HFF cells in 12 well plates. Parasite burden is represented as plaque forming units (PFUs) per organ. Error bars represent mean ± SEM. Data are representative of 2 independent experiments, n=4.

185 Figure 4.8. SRS57-mediated protection is dependent on complement but is not mediated by serum killing of parasites. (A) Survival curve of C57BL/6 mice (Jackson) and C3-/- mice (Jackson) infected with 2x103 CZ1 tachyzoites intraperitoneally. C3-/- mice died significantly sooner than wild type mice (p = 0.041). The results are an aggregate of two independent experiments, 9 mice per group. (B) Survival curve of wild type C57BL/6 and C3-/- mice infected with 2x103 CZ1 Δsrs57 tachyzoites intraperitoneally. No significant difference (ns) was observed between the two strains of mice. The results are representative of one experiment, n=3 C3-/-, n=4. (C) CZ1 and Δsrs57 parasites incubated in 40% NHS for 30 minutes at 37°C. Parasite viability was measured by flow cytometry with a fixable viability stain (Life Technologies) (D) membrane attack complex formation using monoclonal mouse α-human C5b-9 (1:500, Santa Cruz Biotechnologies)

186

Figure 4.9. SRS57 alters systemic and local inflammatory cytokine production. 6-8- week-old C57BL6 mice were infected with 2x103 CZ1 and Δsrs57 tachyzoites intraperitoneally and sacrificed 6 days post infection. (A) Sera and (B) supernatants from ex vivo splenocytes that were cultured for 24 were collected at 6 days post infection for measuring IL-12p40, IFN-g, IL-6, TNF-a, and IL-10 by ELISA (R&D Systems). Data are representative of 2 independent experiments, n=4. Data are represented as mean ±SEM and p-values were obtained using Student’s t-test, *p<0.05, **p<0.01, ***p<0.001.

187

Supplemental Figure 4.1: Schematic representation of the strategy for △srs57 complementation and expression of GFP-LUC. 107 RH Dsrs57 and French RH Dsrs57 (CL55) parasites were co-transfected with a 3.7 kb PCR amplified genomic locus of TgSRS57, including the promoter and the 5’ and 3’ untranslated regions plus a pGFP-LUC plasmid, containing cDNA sequences of the green fluorescence protein (GFP) and firefly luciferase protein (LUC) for in vivo imaging experiments. ~10 days post-transfection, stable parasites expressing GFP were FACS sorted onto 96 well plates. Positive clones expressing GFP were analyzed for TgSRS57 surface expression by flow cytometry. The generation of the complemented strain was done by Dr. Viviana Pszenny, NIH

Supplemental Figure 4.2: Extracellular survival of French strains. 103 tachyzoites of each strain (French RH WT, French Δsrs57 and complement) were maintained for 0 min, 30 min, 1, 2, 4 and 8 hrs. in extracellular conditions, before applying to a fresh monolayer of HFF cells. After 5 days, the number of plaques formed at each time point were counted. No significant differences were observed in the extracellular survival and plating efficiency between the strains. Values represent the mean of triplicates ± SD.

188

Supplemental Figure 4.3. Paired comparison of extracellular survival between RH WT and French RH WT strains. 103 tachyzoites of each strain (RH WT, Δsrs57 and complement and French versions) were maintained for 0 min, 30 min, 1, 2, 4 and 8 hrs in extracellular conditions, before applying to a fresh monolayer of HFF cells. After 5 days, the number of plaques formed at each time point were counted. Linear regression analysis of a paired comparison between the slopes of RH versus French RH strains: (A) wild type, (B) knockout, and (C) complemented, was performed. No significant differences were observed regardless of the strain investigated. Values represent the mean of triplicates ± SD.

189 CHAPTER 5: DISUSSION

5.1 Importance of distinct yet cooperative roles for SRS proteins in regulating the human complement system

Very little is known about how Toxoplasma gondii protects itself from immune defenses during the early stages of infection. Previous work determined that Toxoplasma gondii was resistant to serum killing and suggested that there are several parasite surface proteins that are C3b targets (173). However, it has been 30 years since this work was done and we still do not know the parasite factors responsible for mediating serum resistance or the biological role of complement during T. gondii infection in vivo. The work in this dissertation adds to the scarce body of T. gondii literature about complement activation by identifying the parasite factors targeted by C3b, providing significant insight into the activating nature of the parasite surface and a mechanism for parasite resistance to serum killing. In Chapter 2, I identified significant differences in the level of C3b deposited on the surface of Type I and Type II strains. This observation allowed us to use a forward genetics approach to identify parasite genes relevant to serum resistance and we showed that complement deposition is a complex trait involving several parasite factors. I identified two surface coat proteins, SRS29C and SRS25, that were associated with C3b deposition, confirming our hypothesis that SRS proteins were significantly associated with complement regulation. In Chapter 4, using a reverse genetic strategy, we independently identified SRS57 as another potentially universal C3b acceptor that promoted C3b deposition on both Type I and Type II strains. By engineering parasites deficient in these

SRS proteins, I determined they exhibited distinct yet cooperative roles in interacting with

190 and regulating both lectin and alternative pathways of the complement system. Most notably, using these strategies, I showed that T. gondii SRS proteins acquire host regulatory proteins Factor H and C4b-binding proteins to mediate serum resistance, and thus assigned novel biological functions to a family of proteins whose biological function was poorly understood.

Since this is the first study to characterize complement activation and regulation on the T. gondii surface by SRS surface proteins, a major goal of this work was to gain a better understanding of the precise factors that facilitate SRS-mediated serum resistance. SRS proteins share 25-80% sequence identity, so it is perhaps not surprising that these proteins have redundant functions, as demonstrated by the capacity of both SRS29C and SRS25 to recruit FH and C4BP. Despite this redundancy, these proteins regulate C3b deposition in distinct ways. It is known that the SRS proteins are also polymorphic and exhibit strain specific differences in their expression (260, 261), however the specific contribution that polymorphism has on their biological function is not known. Thus, we tried to distinguish what factors contribute to complement activation versus those that had regulatory roles for these proteins.

The differential expression of SRS29C was one important factor that contributed to the differences in levels of C3b deposition between Type I and Type II stains. Although our QTL was based on polymorphism and not expression, it is conceivable that polymorphisms in the SRS29C 5’UTR likely contribute to expression level differences between Type I and Type II strains. An important finding of this dissertation work was that

T. gondii activated the lectin pathway, which suggested that T. gondii carbohydrate ligands were specifically recognized by MBL and ficolins. Our data indicated that MBL binding

191 was SRS29C-dependent and provided novel insight into the role N-glycosylation of T. gondii SRS29C has in promoting innate immune recognition. Interestingly, one non- synonymous SNP affected a predicted N-linked glycosylation site at amino acid position

113 (serine to asparagine), though the significance of this polymorphism in MBL binding was not investigated. However, using an SRS29C transgenic over expressing strain, we did observe interesting differences in carbohydrate ligands that could provide clues. A flow cytometry based lectin binding assay to characterize N-linked carbohydrate ligands on

Type I RH, Type II Me49 and an RH strain transgenically over expressing the Type II

SRS29C allele showed that lectins ConA, WGA, and RCA bound less well to Type I strains, implying that they had less mannose, GalNAc and GlcNAc present on their surface.

Although SRS29C++ parasites exhibited similar expression levels of SRS29C as Type II strains, the binding of lectins was not completely restored (Sikorski, unpublished observations). Although SRS29C++ parasites exhibited similar Type II levels of ConA binding, they did not demonstrate increased binding of the other lectins specific for

GlcNAc and GalNAc (Sikorski, unpublished observations). Similar results were obtained for a Type I Dsrs29B Dsrs34A double knockout strain that naturally over expresses SRS29C

(261), providing further evidence for distinct differences in N-glycan modifications of

SRS29C between Type I and Type II strains. It important to note that although SRS29C++

Type I strains exhibited significant increases in C3b deposition, it was still 10-fold lower than what is observed for Type II strains, indicating that the expression level of SRS29C is not the only factor that could be controlling C3b deposition. Mutagenesis of SRS29C- specfic N-glycosylation sites would more formally assign a role for surface N-glycans in

MBL recognition and their contributions to lectin pathway activation. Furthermore, results

192 from the lectin binding assay indicated that there are additional unidentified glycosylated surface proteins on Type II strains over and above that accounted for by SRS29C expression. Based on this data, we speculate that the “unaccounted-for” glycosylated surface proteins likely exist as additional factors contributing to complement recognition and activation. Such studies would assign a potential role for glycosylation of T. gondii surface proteins in innate immune recognition.

In Chapter 4, we independently identified SRS57 as another potential C3b acceptor using reverse genetics. It was odd that SRS57 was not identified in the QTL analysis, as it is highly polymorphic between Type I and II strains. However, no expression level differences exist between Type I and Type II strains, suggesting that SRS57 surface expression may be more relevant than polymorphism for its ability to bind C3b, which is certainly a testable hypothesis by the transgenic over-expression of either allele in a ∆srs57 mutant strain. Unlike SRS29C, the complement activating nature of SRS57 is not clear.

SRS57 has no predicted N-linked glycosylation sites (NetNGlyc1.0), however it is predicted to contain several O-linked glycosylation sites (NetOGlyc4.0), suggesting that it may contain carbohydrate ligands that can activate the lectin pathway. Previous work has determined that T. gondii has the capacity to post transitionally modify proteins through

O-linked glycosylation (495). The lectin binding assay employed in Chapter 2 was specific for N-linked sugars and could not be used to test if the decrease in C3b deposition on

Dsrs57 parasites correlated with a loss of O-glycans. Unfortunately, there are few commercially available lectins that recognize complex O-linked sugars and a lack of a universal enzyme to cleave O-glycans from protein backbones, encumbering the possible study O-glycans experimentally (496). However, plant lectin Vicia villosa has recently

193 been used to identify O-linked T. gondii proteins, one of which was SRS44, a tissue cyst wall protein (497). It is possible to use a similar lectin affinity chromatography approach to determine if SRS57 possess an O-linked GalNAc. It is likely that ficolins may play a greater role in recognizing acetylated structures associated with O-linked glycosylation

(498), thus flow cytometric assays could help determine if MBL or ficolins recognize

SRS57 and facilitate complement activation.

Unlike SRS29C or SRS57, we determined SRS25 limited C3b deposition in both

Type I and Type II strains and did not apparently function as a C3b acceptor. However, we do not have enough evidence to completely rule out SRS25 as an acceptor. Because antibodies against SRS25 are lacking, our capacity to further characterize this protein has been limited. SRS25 had fewer SNPs between Type I and Type II strains than SRS29C and it is not known if there are any expression level differences between Type I and Type II strains, however there are also several SNPs in the 5’UTR region. Thus, it remains unknown whether allelic differences in SRS25 contribute to the differential C3b deposition between Type I and Type II strains, or whether this is simply a reflection of the differences in SRS29C expression between these two strains. Considering our data demonstrates that

SRS25 mediates the majority of FH recruitment, one cannot rule out that polymorphism may impact FH binding and thereby alter the levels of C3b that gets surface deposited.

Considering that SRS25 is relatively conserved and expressed in all parasite stages, it is likely that it plays a similar role in all parasite strains and stages of the parasite life cycle in limiting complement activation. Further studies are needed to determine if there are species-specific and stage-specific differences in the ability of SRS25 to bind FH and limit

C3b deposition.

194 In this dissertation, I show that FH down regulates the amplification of the AP on the parasite surface and plays a critical role in protecting parasites from serum killing.

Another major goal of this study was to determine how parasites acquire FH. An interesting finding was that SRS29C, SRS25, and SRS57 all bound Factor H to varying degrees. This is not surprising in the case of SRS29C and SRS57, considering that FH must be in the vicinity of C3b to mediate its regulatory functions. This also suggested that these proteins exhibit analogous properties or employ similar mechanisms to acquire FH. Several studies have shown that bacterial surface proteins can form tripartite complexes with C3b and FH

(499–501), however the molecular or structural basis for these specific interactions with

FH have not all been determined. One study recently proposed that FH possesses a common microbial binding site for several pathogens that coincides with its heparin binding site domains 19-20 (477). The best characterized structural analyses have determined that the discriminatory capacity of FH to bind to C3b coated host cells requires the dual recognition of C3b and host sulfated proteoglycans (130). Thus, I hypothesized that T. gondii’s lectin- specific activity for host sulfated proteoglycans, predicted to be mediated by SRS proteins, was a mechanism for FH recruitment to the parasite surface.

In order to gain insight into the structural basis for recruitment of FH by SRS proteins, we solved the crystal structures for SRS29C and SRS57 in collaboration with Dr.

Martin Boulanger from the University of Victoria. Although our results indicated that

SRS25 was the major recruiter of FH, SRS29C and SRS57 fulfilled both requirements for

AP-mediated regulation by their ability to bind both C3b and FH. In Chapter 3, the crystal structure of SRS29C demonstrated a dimeric configuration with a deep basic groove capable of docking a host sulfated proteoglycan. We engineered a Type I strain

195 transgenically overexpressing SRS29C with mutations to abrogate the basic charge within the SRS29C homodimer groove and showed that a loss in SPG binding also abrogated FH recruitment, supporting our hypothesis. This mechanism is significant because many microbial pathogens acquire FH through its heparin binding domains, suggesting SPG could be a critical mediator of this interaction as an alternative interpretation of those findings.

We next addressed the predictions made by comparative modeling studies based on the crystal structure of SRS29B that indicated that SRS57 exhibited a deeper basic groove that could also accommodate host polyanions like sulfated proteoglycans (265). In Chapter

4, contrary to previous predictions (265), we reported that the crystal structure of SRS57 did not dimerize nor did it possess a basic groove. These results suggested that SRS57 must utilize an alternative mechanism to recruit FH. One possibility is that SRS57 may have an

SPG-independent mechanism for binding FH directly, as suggested for several known pathogens that bind FH through a common site (477). To test this, binding assays between

SRS57 and recombinantly expressed FH that has mutations in the common binding site

(domains 19-20) would be required. A second possibility is that SRS57 forms a heterodimer complex with another SRS protein or complexes with a secreted factor that binds SPG (408) to facilitate C3b inactivation. Recently, large complexes of Plasmodium merozoite surface proteins and 6-CYS proteins have been identified (474, 502), suggesting this could also be a possibility for T. gondii surface proteins. Recent studies have shown T. gondii secreted effector proteins form complexes to synergistically evade innate immunity.

ROP5, ROP17, and ROP18 were found to form heterogenous complexes to collectively block the IRG pathway in mice to synergistically affect acute virulence (503). These

196 studies lend support to the idea that T. gondii SRS proteins may similarly exert their functions synergistically to regulate the complement system by forming complexes, however experimental evidence for SRS heterodimerization is lacking. Further biochemical and structural analyses would determine if this is a feasible hypothesis.

Future studies should include immunoprecipitation of SRS57 in the presence of serum to determine if SRS57 complexes with other SRS proteins and FH in order to better elucidate the mechanism for FH recruitment.

We are attempting to crystallize SRS25 to determine if it acts similarly to SRS29C in binding SPGs or supports FH recruitment independent of SPG binding. Considering the cooperative role of C4BP in serum resistance, identifying the mechanism for its recruitment would also provide a better understanding for how this protein is acquired by T. gondii.

One study has indicated that it also has heparin binding domains, suggesting that it could use similar mechanisms for recruitment to host cells (131). The same SRS29C transgenic strains investigated in Chapter 3 could be used to determine if C4BP is also recruited in an

SPG-dependent manner.

From these findings, it is apparent that conserved proteins, strain-specific differences in SRS protein expression levels, and post translational modification of SRS proteins all contribute to C3b deposition. It is also clear that these proteins are somewhat redundant in their interactions with C3b and FH. Importantly, the apparently distinct SPG- dependent and -independent mechanisms that promote FH recruitment may indicate that these interactions rely on multiple players. It is unclear if the few polymorphisms or post translational modifications in the above SRS proteins or if they contribute to the array of affinities observed for FH and C4BP among the SRS proteins and contribute to strain-

197 specific differences in C3b deposition. In support of this possibility, one study investigating the effect of polymorphism in the gene encoding S. pneumoniae FH binding surface protein

CbpA found that that allelic differences in this gene impacted human FH binding (401).

Based on the findings described in Chapter 2, and from those described above, we developed a hypothetical model for the cooperative regulation of the complement system by T. gondii SRS proteins, depicted in Figure 5.1. We propose that lectin pathway activation is initiated by the detection of N-glycans associated with SRS29C, and possibly through O-linked sugars predicted to be present on SRS57, resulting in the formation of

LP C3 convertases (C2aC4b) which activate C3. SRS29C and SRS57 is then covalently modified by C3b. C4BP is recruited by SRS29C through unknown mechanisms and mediates co-factor activity for Factor I to inactivate C4b and inhibit further C3 and C5 convertase formation. However, our data indicate that C4BP alone does not contribute greatly to parasite resistance, suggesting it is less efficient at inactivating C3b (108). Thus,

Factor H is recruited by SRS29C through its interaction with SPG, which likely increases its affinity for C3b. Co-factor activity of FH facilitates the inactivation of C3b to iC3b by

Factor I and prevents C3b from forming AP C3 convertases (C3bBb). SRS57 and SRS25 also recruit FH through either an SPG-dependent or -independent mechanism to inactivate

C3b and prevent AP C3 convertase formation to suppress the C3b amplification loop. It is possible that SRS57 and SRS25 form a complex either with each other or with one of the other 10 immunogenic SPG-binding proteins identified in this study to facilitate FH recruitment. Together, SRS29C and SRS25 work cooperatively to promote C3b inactivation and prevent the cascade from progressing to the terminal pathway and thereby inhibit formation of the membrane attack complex to ultimately protect the parasite from

198 complement-mediated lysis. To our knowledge, this is the first report identifying the T. gondii factors responsible for mediating serum resistance. The role of SRS57 and its cooperativity with SRS29C and SRS25 in serum resistance is not known, and would require the generation of double (Dsrs29CDsrs57 DKO, Dsrs25Dsrs57 DKO strains) and perhaps even triple knockout (Dsrs25Dsrs29CDsrs57 TKO) to deduce its contribution to complement activation and serum resistance.

5.2 Implications for the use of the murine model for complement studies

Sufficient complement activity is one factor that is important to its protective effect, however over induction of this response can also result in deleterious effects contributing to inflammatory disease. Several factors are known to impact complement activity, including age, sex and species (396, 397, 504). In both humans and mice, females exhibit much lower levels of C3 and terminal pathway components (397, 504). Polymorphisms in complement genes could also impact complement levels and activity (505–507), which have been associated with increased disease risk for several infectious diseases, including susceptibility to intracellular protozoan pathogens, including Cryptosporidium,

Leishmania and Plasmodium (505, 508–512). Interestingly, case studies of two individuals with hereditary C6 deficiency and one patient with terminal complement deficiency reported that these individuals presented with toxoplasmosis (513–515), indicating that complement may play an important role in providing protection against T. gondii in humans. Several studies in the malaria field have found that polymorphisms in the MBL gene may affect the severity of Plasmodium falciparum infection, indicating that a higher percentage of patients with severe malaria had lower levels of MBL associated with mutations in the MBL2 gene (509, 510). When I tested serum from a patient with a

199 mutation in the MBL gene that resulted in loss of MBL function, I found a subsequent reduction in complement deposition in response to T. gondii, so it is likely that mutations in the MBL genes affect functionality or serum levels and also contribute to differences in complement activation in the human population infected with T. gondii. Several polymorphisms in the FH gene could conceivably also affect both C3b and SPG binding and be associated with human disease (516, 517), thus it would be intriguing to test serum from several of these individuals to determine if this impacts FH recruitment to the parasite surface. Whether polymorphisms in complement genes affect complement activation or if complement binding to the parasite surface alters the pathogenesis or disease outcome are not known.

Experimental mouse models have been used extensively to understand the role of complement in infection and disease. The acute susceptibility of the Dsrs29CDsrs25 DKO parasites in NHS suggests the SRS29C-C4BP and SRS25-FH interactions could be attractive targets to block therapeutically, or as possible vaccine targets that could increase the parasite’s susceptibility to host defenses. However, to study any complement- enhancing therapeutic effects would require a suitable in vivo model. In Chapter 2, we determined that mouse and human complement are vastly different in their complement activities. We showed that Dsrs29CDsrs25 DKO parasites exhibited little lysis in 10% fresh mouse serum, and required up to 80% mouse serum to achieve ~60% lysis in 30 minutes

(unpublished observations), suggesting that mice are less efficient at killing parasites present in non-immune serum. Indeed, no differences in survival was observed when the

Dsrs29CDsrs25 DKO parasites were injected intraperitoneally, indicating the mouse model may not be suitable for studying the lytic function of complement in vivo.

200 Similar observations using the mouse model were made in studies investigating the role of surface lipophosphoglycan LPG in Leishmania infection. LPG is a major surface component of metacyclic Leishmania parasites that has been implicated for the establishment of infection, including complement evasion, attachment and entry into macrophages (518). However, LPG deficient parasites (lpg1-) that were sensitive to lysis in as little as 1% human serum did not lyse in incubations with up to 60% fresh mouse serum (406). Furthermore, no differences in pathogenesis was observed when lpg1- parasites were infected in either wild type or C5 deficient mice, indicating that complement mediated lysis plays no role in the mouse model either in vitro or in vivo (406). A recent study indicated that mouse serum contains complement inhibitors that strongly inhibit classical pathway activation (519), suggesting that even studies investigating targeted antibody-based strategies to increase complement activity may not be successful. However, in Chapters 3 and 4, we determined that the mouse model is still useful in studying other aspects of complement effector functions. Regardless of the inefficiency of complement mediated lysis in non-immune mice, C3 deficient mice were more susceptible to acute T. gondii infection, establishing a protective role for complement and indicating that effector functions outside of direct killing by complement may be of more importance.

The differences observed between human and mouse complement brings up interesting questions about the coevolution of T. gondii with various hosts, species-specific differences in binding affinities for complement regulator proteins and the degree to which they could contribute to parasite resistance. Factor H is known to exhibit sequence variations among mammalian species, with only 60% sequence homology between human and mouse FH (401). Because lysis of the Dsrs25Dsrs29C DKO was observed in 80%

201 mouse serum, it suggests that murine FH recruitment may contribute to parasite resistance to serum killing. However, even in high concentrations of mouse serum, the DKO parasites were much less susceptible to killing compared to human serum, perhaps suggesting that

SRS29C and SRS25 may not be the relevant FH binding proteins that regulate mouse complement. Alternatively, it is possible that SRS57, or another dominant SRS surface protein, may exhibit higher affinity for murine FH and could play a larger role in mediating protection in the murine host. Several studies provide support for this hypothesis by demonstrating that several established pathogen FH binding proteins that mediate resistance against human complement have little or no affinity for murine Factor H and thus were dispensable for protection in the murine model (401, 520, 521). To address these questions, similar flow cytometric assays using mouse serum to measure binding of murine

FH and C4BP in the context of SRS29C, SRS57, and SRS25 could provide answers to these questions. Regardless of these species-specific differences in complement activity, the complement system has evolved to overcome pathogen subversion of the lytic response by employing several other effector functions. Considering T. gondii is resistant to both human and mouse serum, the study of additional complement effector functions may be more relevant and would promote better understanding of the protective effects of the complement system that extend beyond complement-mediated lysis.

5.3 Defining novel C3-dependent biological functions of SRS29C and SRS57 in regulating parasite virulence

Virulence is a complex phenotype involving several parasite and host factors. The parasite must balance immune evasion with immune activation in order to establish a chronic infection to ensure a transmissible infection. Identifying SRS29C and SRS57 as

202 C3 acceptors on the surface of T. gondii is an important finding because C3 is a potent immunological ligand for complement receptors, and this ability to sense parasite infection may well contribute to the induction of a measured immune response that regulates parasite proliferation without inducing significant immunopathology, striking the correct balance that permits both host and parasite survival. In Chapters 3 and 4, the biological significance of these interactions was evaluated in vivo. Until now, the biological role of SRS proteins and the host factors they interact with to mediate their function was poorly understood. The in vivo studies presented in this dissertation demonstrate that SRS29C and SRS57 mediate their biological effects by interacting with complement.

Using the murine model, we established that both SRS29C and SRS57 attenuate virulence to prolong host survival and promote parasite persistence. In Chapters 3 and 4, both Dsrs29C and Dsrs57 parasites exhibited increased virulence, and mice experienced elevated IL-12 and IFN-g production and succumbed to infection more rapidly than mice infected with wild type parasites. We also showed that infection of C3-/- and WT mice with

Dsrs29C or Dsrs57 parasites had no difference in survival, indicating that SRS-mediated attenuation of virulence was C3-dependent. However, unlike Dsrs29C infected mice,

Dsrs57 infected mice additionally exhibited increased parasite burden 6 days post infection, suggesting that the dysregulated cytokine production could be simply the result of uncontrolled parasite expansion. This was not the case for SRS29C, where no difference in parasite burden between Dsrs29C and WT parasites was observed at day 6 post infection, indicating that these differences in pathogenesis could be immunologically driven. Mice infected with RH SRS29C++ parasites exhibited prolonged survival compared to wild type strains and a reduction in systemic IL-12 and IFN-g production early in infection that was

203 not dependent on parasite burden. This is significant because mice infected with lethal

Type I strains die by an over exuberant Th1 response that is referred to as a cytokine storm

(342). These data suggest that SRS29C over expression induces a more controlled immune response early during acute infection. Differences in parasite burden were observed only after 7 days post-infection, suggesting that early alterations in immune induction contributed to the significant decrease in parasite burden. To further support this deduction, co-infection assays with wild type RH stain and the RH strain transgenically over expressing SRS29C showed that SRS29C expression was able to control the proliferation of WT parasites in trans and protected mice from lethal infection, indicating that this protection was immune driven. To further confirm these findings, RAG2-/- or SCID mice, deficient in adaptive immunity, could be assayed to establish whether parasites expand equally in the absence of an adaptive immune response.

Considering our in vitro findings that demonstrated both Dsrs29C and Dsrs57 parasites exhibit reduced C3b deposition, it is unclear why differences in parasite expansion existed. It is possible these parasites experience different growth kinetics in vivo.

Similar co-infection assays could be performed between WT and Dsrs57 parasites to determine if SRS57 expression regulates parasite proliferation in trans. Given that SRS proteins appear to interact with multiple host factors, it is possible that SRS57 is also recognized by another receptor on host cells that works synergistically to control parasite proliferation. For example, several studies have shown that iC3b recognition through CR3 works cooperatively with several other receptors, such as TLRs, CD14 and Dectin-1, to amplify or inhibit immune cell signaling pathways that regulate immune responses, as described for several pathogens (486, 487, 522).

204 While these results established that SRS29C and SRS57 mediated protection in vivo was dependent on their interactions with C3, the complement-mediated mechanism driving this phenotype was not determined in this dissertation. Since the predominant form of C3b on wild type parasites was iC3b, we hypothesized that FH mediated inactivation of C3b contributed to our phenotype. In support of this, in vivo studies comparing wild type RH and strains overexpressing wild type SRS29 and the K16/17S mutated version showed the

SRS29C-mediated protection was lost as a result of the mutation in the SPG-binding site that abrogated FH recruitment. Although SRS29C++ and SRS29C++ K16/17S parasites had equivalent levels of C3b deposition, western blot analysis showed the K16/17S mutation exhibited reduced iC3b levels. We reasoned, based on these results, that the interaction with FH and its C3b regulatory capacity was an important factor driving this in vivo response. However, the contribution of FH mediated-downregulation of complement to pathogenesis in vivo was never directly shown. FH deficient mice were considered to test this directly, however this approach may not have resolved this phenotype. FH deficient mice exhibit rapid C3 consumption due to uncontrolled complement activation and succumb to autoimmune disease (523), thus using these mice could lead to misinterpretation of the results. The authors of a recent study evaluating the role of FH recruitment by Bacillus anthracis surface pore protein BclA in promoting spore persistence rationalized using the C3 deficient model for similar reasons (524).

5.4 Implications for complement-mediated lysis independent mechanisms of host protection during acute T. gondii infection and future directions

A host protective role for complement seems counter intuitive in the context of complement-resistant parasites, as one would expect more resistant parasites to be more

205 virulent. Furthermore, Dsrs29C and Dsrs57 parasites should theoretically be less virulent in the context of reduced FH recruitment. However, despite reductions in FH recruitment, these parasites are still resistant to complement-mediated lysis due to the redundant recruitment of FH by SRS25. Thus, the phenotypes observed in vivo must be decoupled from complement-mediated lysis in vitro, as mouse complement is not lytic in vitro, and it appears to be dispensable for murine protection in vivo. We reasoned that the loss of the host protective effect in C3 deficient mice could be attributed to the changes in the levels and activation of C3b on the parasite surface in the absence of SRS29C and SRS57, resulting in alterations in one or more downstream complement effector functions which include clearance by phagocytosis, stimulation of inflammatory responses or stimulation of adaptive immunity. One limitation in using C3 deficient mice is that this model cannot distinguish between two potential effector proteins, C3b or C3a, and their relative and contributing roles in mediating protection in vivo. The responding population of immune cells that recognize these effectors and control the infection are also critical factors in this response, thus the cellular basis for complement-mediated host protection is another question that remains unanswered.

In this section, I will discuss how SRS-mediated alterations in complement activation could impact several complement effector functions that may play a role in regulating parasite proliferation and the inflammatory responses. Complement activation is a potent inflammatory response that must be regulated to prevent damaging inflammation in the host. In this case, T. gondii may exploit its interactions with complement to promote its survival by capitalizing on its link with inflammatory responses or adaptive immunity. In Chapter 2, we determined that T. gondii is opsonized with

206 inactivated complement protein iC3b. In Chapter 3, we used iC3b-coated parasites to determine if opsonization impacted cytokine production in vitro. We infected bone marrow derived macrophages (BMDMs) with WT Type II parasites that were incubated with 40%

NHS and washed to remove the influence of anaphylatoxins (C3a and C5a) and their interaction with cognate receptors C3aR and C5aR. Using a Luminex assay, we observed reduced production of cytokines IL-6 and chemokines MIP-1a, MIP1-b, Mip2, KC, and

RANTES production compared to BMDMs infected with parasites that were not previously exposed to serum. There were no differences in the infectivity of the parasites, thus these findings suggest that parasite regulation of complement activation on the parasite surface can alter cytokine and chemokine production, possibly through iC3b interactions with complement receptors CR3 and CR4. These results are in line with several studies that have demonstrated the binding of inactivated complement (iC3b) to complement receptor CR3 which suppressed inflammatory cytokine production (428, 429, 525, 526). Furthermore, several pathogens, including Leishmania, exploit opsonization with iC3b to gain entry into host cells to promote parasite survival (527, 528) Based on these results, one could speculate that parasites exhibiting less iC3b, or no C3 in the context of C3 deficiency, or parasites in the absence of serum, may be less able to down regulate inflammatory cytokine production via iC3b-mediated interactions with complement receptors. We can also not rule out the effect of other complement proteins, thus using C3 deficient serum would establish that this was a C3 mediated effect. From this in vitro assay, however, we could not determine if the parasites entered the cells through invasion or complement-mediated phagocytosis to impact this response. Until recently, there were only limited tools available to distinguish between invasion and phagocytosis using flow cytometry. Recent studies

207 have developed a flow cytometry based assay exploiting pH sensitivities of fluorescent markers to distinguish phagocytosis from invasion during T. gondii infection (529).

Utilizing such strategies provides a way forward to study the role of complement opsonization on both in vitro and in vivo phenotypes.

An important factor to take into consideration is the efficiency of complement- mediated phagocytosis in clearing parasites, in order to determine if this response is protective. While the primary mode of entry for T. gondii into host cells occurs via active invasion, complement-mediated phagocytosis cannot be ruled out. Several studies have shown the Fc-mediated endocytic uptake of antibody-opsonized parasites results in parasite killing (266, 415). In general, complement-mediated phagocytosis is distinct from Fc- receptor mediated phagocytosis in that it is not particularly efficient in internalization, does not induce respiratory burst, and requires priming with IFN-g to be more efficient (82, 530–

533). However, studies investigating the role of complement opsonized T. gondii in phagocytosis are lacking.

There are several factors that contribute to efficient opsonin-mediated phagocytosis that could be exploited by T. gondii to alter immune outcome. First, the density of ligands, such as iC3b or C3b, on the phagocytic target is a critical factor in the interaction with phagocyte receptors, where lower densities of IgG and iC3b have been associated with reduced capacity of the phagocyte to bind its target (534). Binding of multiple ligands and aggregation, also known as crosslinking, of phagocytic receptors increase affinity for its ligand and promotes activation of these receptors to facilitate endocytosis (535). This perspective could provide insight into the biological relevance of the significant differences in C3 deposition between Type I and Type II strains. Considering there is evidence that

208 Type II strains preferentially enter murine macrophages via phagocytosis to invade from within the phagosome (308), perhaps the increased complement deposition on Type II strains prolongs or strengthens the affinity for interactions with phagocytic receptors like

CR3 and CR4, which promote a preferential uptake of Type II strains. Conversely, the low levels of C3b on Type I strains may be a function of too few ligands to promote sustained interactions with host cells, thus favoring active invasion over endocytosis. Second, this interaction often requires additional stimuli through additional receptors, including TLRs

(536) or CD44, to increase the affinity of CR3 for its ligand. It is plausible SRS29C binding of SPGs and, as discussed above, potential interactions of SRS57 with additional receptors, could provide additional signals that alter the immune response. Recent studies investigating the lectin activity of T. gondii microneme proteins MIC1 and MIC4 has shown that they bind TLR2 and TLR4, providing the potential for cooperative signaling between several T. gondii proteins and host cell receptors to regulate innate immune priming (537).

Evaluating the role of CR3 and CR4 in complement-mediated phagocytosis in vivo may not be straightforward. CR3 and CR4 bind ligands that contribute to their diverse functions. For example, CR3 possesses a lectin domain which recognizes pathogen- associated molecules such as b-glucan (538–540). It is also an integrin that binds endogenous ligands to promote adhesion and recruitment. Previous work showed that blocking CR3 during acute T. gondii infection significantly increased susceptibility to infection with avirulent Type II Me49 parasites(541). The authors determined that mice died of severe necrosis and inflammation, however they also observed significant reductions in the amount of polymorphonuclear leukocytes, CD4+ T cells, and CD8+ T

209 cells in the peritoneal cavities of these mice. However, due to its multiple functions, they could not rule out that blocking CR3 had not impacted its integrin function. Therefore, the reduced capacity to recruit cells to the site of infection may have contributed to uncontrolled parasite replication and mortality. Hence, infection of CD11b/CD18 (CR3) deficient animals to determine the role of CR3 would likely bring similar complications in interpreting phenotypes and would not resolve the question. Knock down or blocking of

CR3 in murine BMDMs or BMDCs in vitro may provide more definitive answers in determining whether CR3 plays a role in complement-mediate phagocytosis or alterations in cytokine production.

CR2 is also an important complement receptor on B cells that recognizes iC3b and

C3dg bound antigen and is critical for stimulating B cell responses. As antigens bound by iC3b and C3dg, SRS29C and SRS57 may also induce antibody responses. During acute infection, SRS-specific antibody responses are thought to play a critical role in targeting the adaptive immune response towards tachyzoites to limit parasite proliferation (259, 268,

364, 368). During acute infection, SRS29B and SRS34A are two highly immunogenic tachyzoite proteins that dominate the early humoral response. In our early studies, we observed that immune serum from mice infected with strains that naturally express high levels of SRS29C, such as Type II and Type III strains, as well as those infected with RH

SRS29C++ possessed strong antibody responses against SRS29C. Hence, the presence of antibodies against SRS29C detected in acute infection serum from mice 8 days post- infection correlated with SRS29C expression, and infection serum from mice with low

SRS29C expression, such as RH, failed to produce these antibodies (unpublished observations). This suggests that higher levels of complement opsonization could be one

210 mechanism driving SRS29C specific humoral responses. However, SRS57, which is highly expressed on all strains, is also tagged with iC3b, but does not produce an immunodominant response. If this is a complement dependent response, it is possible that

SRS57 may not be a major murine C3 acceptor, thus explaining its marginal contribution to simulating a humoral response. Further studies characterizing the interactions of these

SRS proteins with mouse complement would be necessary to confirm these suppositions.

Alternatively, complement is not the only factor that determines immunogenicity of an antigen and it is unknown what additional differential factors between these two SRS proteins may contribute to this response. One known difference between these two proteins is that SRS29C is glycosylated. Several studies have shown that glycosylation of viral surface antigens influence their immunogenicity (542–544), suggesting that glycosylation is another possible factor that could contribute to the humoral response. Future experiments with C3 deficient mice would need to confirm that this is a C3 dependent phenotype.

The inflammatory nature of the complement system in vivo is attributed predominantly to anaphylatoxins. The increased production of inflammatory cytokines observed in Dsrs29C and Dsrs57 infected mice could alternatively be attributed to changes in the production of anaphylatoxins C3a and C5a. The reduced capacity of FH recruitment to regulate complement activation would theoretically increase the levels of C3a and C5a in the local environment, resulting in chemotaxis of inflammatory cells expressing C3aR and C5aR to the site of infection. To determine if this is indeed occurring in the absence of

SRS29C and SRS57, levels of C3a and C5a could be measured in the serum and in local splenocyte cultures. While reduction in the levels of inflammatory cytokines IL-12 and

IFN-g in the serum of C3-/- infected mice that we observed in Chapter 3 could support this

211 hypothesis, it could not explain the perplexing increase in the production of IL-12 and IFN- g in C3 deficient mice splenocytes. Several studies have reported evidence of a C3 bypass mechanism in C3 deficient mice (427, 545, 546) whereby thrombin-mediated C5 cleavage produces C5b and C5a. We showed that C5a was present in the serum of C3-/- animals infected with T. gondii. C5a exhibits a redundant function in promoting inflammatory responses in mice (152, 427, 547) and is considered to be more potent than C3a, possibly explaining the greater cytokine response of splenocytes from C3-/- infected animals.

Overproduction of C5a contributes to pathology associated with several highly pathogenic viral infections and induces cytokine storms (548, 549). To investigate that this bypass is also occurring locally, assessing ex vivo splenocyte C5a production in C3 deficient animals infected with T. gondii would resolve this question. Moreover, infection of C3aR-/-/

C5aR-/- mice would eliminate this bypass mechanism in order to study the role of anaphylatoxin signaling in acute T. gondii infection.

As T. gondii replicates and disseminates during acute infection, it establishes infection in various tissues. Since serum complement cannot reach all tissues, the question of localized complement production by cell-derived complement is important to consider.

Whether local complement production is activated in situ in response to a local infection has not been assessed in the infection model. Studies evaluating the role of local complement production by immune cells have demonstrated novel, non-traditional functions for complement proteins in regulating Th1 responses, particularly anaphylatoxins, that is independent of serum derived complement. Recent studies have shown that APCs and T cells express C3aR and C5aR and are able to produce their own complement intracellularly upon cognate APC-T cell interactions (152). C3a and C5a also

212 have intrinsic roles in regulating T cells responses that function in autocrine and paracrine fashion to activate T cells and induce Th1 responses. It is also unclear if complement produced by these cells can amplify a localized immune response or if its function is to amplify paracrine signaling through C3aR and C5aR on APCs and T cells to enhance their activation. This is one possible way in which changes in complement activation in response to Dsrs29C and Dsrs57 parasites could indirectly impact T cell responses. One experiment that might lend support to this hypothesis is using C3aR and C5aR antagonists in ex vivo splenocyte cultures taken from Dsrs29C and Dsrs57 infected mice to determine if this reduces IL-12 and IFN-g production to wild type levels. In support of complement regulation as an important factor in regulating T cell responses, DAF-/- mice have been shown to exhibit higher T cell IFN-g production and enhanced T cell responses when stimulated with antigen in vitro (156). This was associated with reduced capacity to regulate C3 convertase activity and increased C3b and C3a production by APCs. Based on these findings, one could speculate FH could play a similar role.

To summarize the findings in this dissertation, we generated a comprehensive synopsis of the transgenic and knock out strains used in these studies and how these genetic changes altered the experimentally identified interactions with complement proteins. We highlight the complement mediated effector functions that could be impacting the immune response to alter pathogenesis and contribute to parasite persistence in vivo (Figure 5.2).

In this dissertation, we established that complement regulation is primarily driven by FH and protection in the murine model is independent of complement-mediate lysis. Thus, the complement-dependent protective effect observed during acute T. gondii infection in the murine model is most likely due to the effector molecules C3b and C3a. We hypothesize

213 that parasites exploit down regulation of complement effector proteins to balance host inflammation. Wild type parasites establish persistent, transmissible infections compared to SRS29C and SRS57 deficient strains or upon infection of complement deficient mice.

Wild type parasites downregulate complement activation, resulting in an increase in iC3b and a corresponding reduction in C3a. Our studies demonstrate that both SRS29C and

SRS57 promote complement deposition and concurrently inactivate C3b through SPG- dependent and -independent recruitment of FH. SRS29C and SRS57 deficiencies thus result in less iC3b, and hypothetically, more anaphylatoxin C3a. Dsrs29C and Dsrs57 infected mice exhibit unregulated inflammation that likely contributes to mortality, suggesting that interactions with complement alter cytokine production to control parasite proliferation and promote persistent infection. SRS25 deletion results in subsequent increased C3b deposition due to loss of FH recruitment. SRS25 SRS29C double deletion reduces both FH and C4BP, increasing levels of active C3b on the parasite surface, and render the parasites susceptible to serum killing.

The complement effector functions mediating this protection remain unknown, however we summarized the possible effector functions in Figure 5.2 that could be play a role in altering cytokine production and the induction of protective immunity. It is possible that loss of FH results in more anaphylatoxin production, leading to the recruitment of immune cells that could further contribute to inflammatory cytokine responses to cause immunopathology. Alternatively, local complement activation by APCs could result in autocrine and paracrine anaphylatoxin production that impacts activation of T cells and

Th1 differentiation. Dysregulation of this response can lead to alterations in Th1 cytokine production. We demonstrated serum coated parasites reduced both cytokine and

214 chemokine production in vitro. At present, it is unclear if CR3 or CR4 mediated phagocytosis plays a role in these alterations and remains to be determined. Proteins tagged with C3b, such as SRS29C or SRS57, could also be recognized by CR2 on B cells to stimulate SRS-specific antibody production. Identifying the cellular basis for these responses provides a way forward for future studies aimed at better understanding the role of complement during acute T. gondii infection.

Taken together, the biological role of the complement system and biological function of SRS proteins during acute T. gondii infection were poorly understood until now. This dissertation has provided critical insight into the distinct contributions of lectin and alternative pathways to the initiations and amplification of the complement response against T. gondii, respectively, and the sophisticated cooperative functions of SRS29C,

SRS25 and SRS57 in suppressing this potent innate immune response by recruiting host regulators C4BP and FH. The structural analyses performed in this dissertation provided mechanistic insight into the diverse strategies in which T. gondii acquires host regulator

FH to its surface, which consist of both SRS29C-SPG-dependent and SRS57-SPG- independent mechanisms. In vivo studies determined that both SRS29C and SRS57 regulated virulence and cytokine production during acute T. gondii infection. Using C3 deficient mice, I showed these mice phenocopied the infection kinetics and altered pathogenesis of wild type mice infected with Dsrs29C and Dsrs57 parasites. Based on this work, the hypothesis going forward is that T. gondii exploits complement regulation to control inflammation and promote parasite persistence. The diverse effector functions of both iC3b and anaphylatoxin proteins C3a and C5a provide opportunities for several lines of investigation for future immunological studies to identify the cellular basis of these

215 responses. Ultimately, this work has assigned several novel biological functions of the SRS superfamily and provided further evidence for their diverse roles in regulating innate immunity to promote the establishment of chronic infection and transmissibility to new hosts.

216 5.5 Figures

Figure 5.1 Hypothetical model of SRS-mediate regulation of the complement system. The lectin pathway (LP) is initiated via mannose binding lectin (MBL) recognition of N- glycans associated with SRS29C and possibly through predicted O-linked glycans present on SRS57. Lectin pathway activation leads to activation of C2 and C4, resulting LP convertase formation which the cleaves C3 into C3a and C3b. C3b covalently modifies SRS29C and SRS57. C4BP is recruited by SRS29C through an unknown mechanism to inactivate C4b into C4d, impeding the formation of LP C3 convertase. FH is recruited by SRS29C in an SPG-dependent manner. Co-factor activity then facilitates C3b inactivation by Factor I to inactivate C3b into iC3b, preventing alternative pathway (AP) C3 convertase

217 formation and LP C5 convertase formation. SRS57 also recruits FH through an SPG- independent mechanism, however the SRS25-mediated mechanism for recruitment of FH is unknown. Together, SRS29C and SRS25 work cooperatively to promote C3b inactivation to regulate the AP amplification loop and prevent the cascade from progressing to the terminal pathway and formation of the lytic membrane attack complexes. Potential hypothetical mechanisms for FH recruitment by SRS57 and SRS25 include direct binding of FH, the formation of SRS heterodimers that can bind SPGs, or interactions with unidentified SPG-binding proteins.

218

Figure 5.2 Synopsis of transgenic and deletion mutants and subsequent alterations in interactions with the complement system. (Top panel) Synopsis of experimentally defined interactions with complement proteins MBL, C3b, iC3b, Factor H, C4BP and host SPGs. Question marks signify lack of experimental evidence and X indicates loss of an interaction due to deletion or no interaction. Both Type I and Type II strains are equally resistant to complement mediated lysis by inactivation of C3b into iC3b, however Type II strains bind significantly higher levels of MBL, FH, and iC3b than Type I strains. MBL recognizes N-glycans associated with SRS29C expression, however post translational modifications associated with SRS57 and SRS25 are unknown. SRS29C and SRS57 promote C3b deposition as C3 acceptors. Deletion of SRS29C results in reduced MBL binding and C3b deposition, whereas overexpression increases iC3b and FH levels on the parasite surface but the impact on MBL binding is unknown. SRS29C binds SPG to recruit

219 FH, which is abrogated as a result of a K16/17S mutation. Deletion of SRS57 also reduces iC3b deposition and FH recruitment but does not bind SPG. SRS25 limits C3b deposition by interacting with FH, thus SRS25 deletion results in subsequent increased C3b deposition. SRS25 SRS29C double deletion reduces both FH and C4BP, increasing levels of active C3b on the parasite surface, and render the parasites susceptible to serum killing. Survival is based on experimental data, where Type I strains are lethal in C57BL6 mice and Type II strains are intermediately virulent. Deletion of SRS29C and SRS57 results in increased virulence, whereas SRS29C over expression in Type I strains significantly prolongs survival. K16/17S mutation abolishes SRS29C-mediated protection in vivo. (Bottom panel) Complement mediated lysis is not a major effector function mediating protection in vivo. Complement effector proteins C3a/C5a and C3b are predicted to mediate protection in vivo. Hypothetical C3a production is based on relative level of C3b deposition. Effector functions include: (left to right) Local production of anaphylatoxins by immune cells to regulate Th1 responses; C3a and C5a mediated recruitment of inflammatory cells to the site of infection; CR3 and CR4 recognition and phagocytosis iC3b opsonized parasites by phagocytes; antigen covalently bound to CR2 lower the threshold for B cell stimulation and antibody production

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