Host-Apicomplexan Parasite Interactions: Leveraging Biological Discovery Into Antiparasitic Drug Development

University of Pennsylvania ScholarlyCommons

Publicly Accessible Penn Dissertations

2013

Host-Apicomplexan Parasite Interactions: Leveraging Biological Discovery into Antiparasitic Drug Development

Melanie Grace Millholland University of Pennsylvania, [email protected]

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Part of the Cell Biology Commons, Molecular Biology Commons, and the Parasitology Commons

Recommended Citation Millholland, Melanie Grace, "Host-Apicomplexan Parasite Interactions: Leveraging Biological Discovery into Antiparasitic Drug Development" (2013). Publicly Accessible Penn Dissertations. 675. https://repository.upenn.edu/edissertations/675

This paper is posted at ScholarlyCommons. https://repository.upenn.edu/edissertations/675 For more information, please contact [email protected]. Host-Apicomplexan Parasite Interactions: Leveraging Biological Discovery into Antiparasitic Drug Development

Abstract The obligate intracellular pathogens Plasmodium falciparum and Toxoplasma gondii remodel their host cell to facilitate their intracellular development and progress through their asexual life cycle, a virulent lytic cycle responsible for parasite-mediated pathogenesis. While several studies have highlighted parasite proteins that interact with the host cell during this cycle, host proteins exploited by the parasite for successful growth and conversely, host molecules evolutionarily tuned to control parasite infection remain unclear. We addressed this question from both sides of the host-parasite interaction in the hope to leverage biological discovery of host molecules involved in infection into the validation of novel drug target candidates with vastly reduced potential for resistance: (1) building on previously established roles of host calpain during the lytic cycle to elucidate host pathways that facilitate parasite life cycle progression and (2) clarifying the existing role of platelets in innate control of parasitic disease. Comparative proteomics, genetics, and biochemical techniques led us to the discovery of a complex Gaq- coupled, protein kinase C (PKC)-mediated host signaling cascade common to both T. gondii and P. falciparum that results in cytoskeletal compromise and facilitates host calpain activation immediately prior to parasite release (Millholland et al., 2011; Millholland et al., 2013). The complexity of this pathway represented an untapped resource of antimalarial targets; indeed, inhibitors of key host signaling components demonstrated antiparasitic activity in murine models of malaria and toxoplasmosis. Conversely, host platelets were previously shown to have antimalarial capacity, leading us to characterize this host-parasite interaction in the hope to harness this antimalarial activity to optimize therapeutic potential. Upon screening molecules secreted in the blood stream, we identified Platelet actF or 4 as the host defense peptide (HDP) component of platelets capable of killing malaria parasites via selective lysis of the parasite digestive vacuole (DV), the site of hemoglobin digestion(Love et al., 2012). To exploit this DV lysis mechanism in a drug discovery effort, we tested a library of small, nonpeptidic mimics of HDPs and identified compounds that potently killed .P falciparum in vitro and reduced parasitemia in a murine malaria model. Taken together, these data reinforce the feasibility of targeting host molecules as a novel antiparasitic strategy.

Degree Type Dissertation

Degree Name Doctor of Philosophy (PhD)

Graduate Group Cell & Molecular Biology

First Advisor Doron C. Greenbaum

Keywords egress, host/parsite interactions, malaria, parasite, platelet factor 4, toxoplasma

Subject Categories Cell Biology | Molecular Biology | Parasitology

This dissertation is available at ScholarlyCommons: https://repository.upenn.edu/edissertations/675 HOST-APICOMPLEXAN PARASITE INTERACTIONS:

LEVERAGING BIOLOGICAL DISCOVERY INTO ANTIPARASITIC DRUG DEVELOPMENT

Melanie G. Millholland

A DISSERTATION

Cell and Molecular Biology

Presented to the Faculties of the University of Pennsylvania

Partial Fulfillment of the Requirements for the

Degree of Doctor of Philosophy

2013

Supervisor of Dissertation

______Doron C. Greenbaum, Ph.D. Assistant Professor of Pharmacology Perelman School of Medicine University of Pennsylvania

Graduate Group Chairperson

______Paul F. Bates, Ph.D. Professor of Microbiology Perelman School of Medicine University of Pennsylvania

Dissertation Committee

James B. Lok, Ph.D. Professor of Pathobiology

Christopher A. Hunter, Ph.D. J. Kevin Foskett, Ph.D. Professor and Chair of Pathobiology Isaac Ott Professor of Physiology

David W. Speicher, Ph.D. Sara Cherry, Ph.D. Caspar Wistar Professor Associate Professor of Microbiology

HOST-APICOMPLEXAN PARASITE INTERACTIONS: LEVERAGING BIOLOGICAL DISCOVERY INTO ANTIPARASITIC DRUG DEVELOPMENT

2013

Melanie Grace Millholland

This work is licensed under the Creative Commons Attribution- NonCommercial-ShareAlike 3.0 License

To view a copy of this license, visit http://creativecommons.org/licenses/by-ny-sa/2.0 ACKNOWLEDGMENTS

Thanks to all collaborators, without whom I could not have completed this work:

Richard Scott, Ph.D. and Katie B. Freeman, Ph.D and everyone at PolyMedix, Inc.;

Photini Sinnis, M.D., Satish Mishra, Ph.D., M. Anna Kowalska, Ph.D., Mortimer Poncz,

M.D., Christopher D. Dupont, Dustin Shilling, Dewight R. Williams, Ph.D. and the following funding sources for this work: NIH R44 AI090762-0; NIHT32AI007532;

University of Pennsylvania TAPITMAT Pilot Program; Penn Genome Frontiers Institute;

Gates Grand Challenges Exploration Program.

Thanks to George Makedonas and Matthew E. Cohen, Ph.D. for helpful scientific and non-scientific discussions.

Thanks especially to my committee members and department chairs for all the open doors to experimental help, unique reagents, listening ears, and always constructive criticism that kept me on track: Sparky Lok, Ph.D., Christopher A. Hunter, Ph.D., J. Kevin

Foskett, Ph.D., David W. Speicher, Ph.D., Sara Cherry, Ph.D., Paul Bates, Ph.D., and

Robert W. Doms, M.D./Ph.D.

Thanks to all current and former members of the Greenbaum laboratory: Melissa

Love, Nataline Meinhardt, Swapnil Kulkarni, Ph.D., Michael Harbut, Ph.D., Bhumit Patel,

Ph.D., Katie St. Denis, Emily Schutsky, Jarrett Remsberg, Eli Berdougo, Ph.D., Hany

Girgis, Ph.D., and Geetha Velmourougane, Ph.D for making me want to come to lab on the dreariest of days.

Special thanks to my advisor, Doron Greenbaum, Ph.D. for giving me enough space to mess up on my own accord, but being the first to celebrate when it finally worked.

iii

ABSTRACT

HOST-APICOMPLEXAN PARASITE INTERACTIONS:

LEVERAGING BIOLOGICAL DISCOVERY INTO ANTIPARASITIC DRUG DEVELOPMENT

Melanie G. Millholland Doron C. Greenbaum

The obligate intracellular pathogens Plasmodium falciparum and Toxoplasma gondii remodel their host cell to facilitate their intracellular development and progress through their asexual life cycle, a virulent lytic cycle responsible for parasite-mediated pathogenesis. While several studies have highlighted parasite proteins that interact with the host cell during this cycle, host proteins exploited by the parasite for successful growth and conversely, host molecules evolutionarily tuned to control parasite infection remain unclear. We addressed this question from both sides of the host-parasite interaction in the hope to leverage biological discovery of host molecules involved in infection into the validation of novel drug target candidates with vastly reduced potential for resistance: (1) building on previously established roles of host calpain during the lytic cycle to elucidate host pathways that facilitate parasite life cycle progression and (2) clarifying the existing role of platelets in innate control of parasitic disease. Comparative proteomics, genetics, and biochemical techniques led us to the discovery of a complex

Gaq-coupled, protein kinase C (PKC)-mediated host signaling cascade common to both T. gondii and P. falciparum that results in cytoskeletal compromise and facilitates host calpain activation immediately prior to parasite release (Millholland et al., 2011; Millholland et al., 2013). The complexity of this pathway represented an untapped resource of antimalarial targets; indeed, inhibitors of key host signaling components demonstrated antiparasitic activity in murine models of malaria and toxoplasmosis. Conversely, host platelets were previously shown to have antimalarial capacity, leading us to characterize this host-parasite interaction in the hope to harness this antimalarial activity to optimize therapeutic potential. Upon screening molecules secreted in the blood stream, we identified Platelet Factor 4 as the host defense peptide (HDP) component of platelets capable of killing malaria parasites via selective lysis of the parasite digestive vacuole (DV), the site of hemoglobin digestion(Love et al., 2012). To exploit this DV lysis mechanism in a drug discovery effort, we tested a library of small, nonpeptidic mimics of HDPs and identified compounds that potently killed P. falciparum in vitro and reduced parasitemia in a murine malaria model. Taken together, these data reinforce the feasibility of targeting host molecules as a novel antiparasitic strategy.

TABLE OF CONTENTS

ACKNOWLEDGMENTS ...... III ABSTRACT ...... IV LIST OF TABLES ...... VI LIST OF ILLUSTRATIONS ...... VI INTRODUCTION ...... 1-10 Apicomplexan Parasite Biology ...... 1 Host-Parasite Interactions Necessary for Survival: Getting In and Getting Out ...... 2 Platelets, Malaria, and Host Defense Peptides: The Other Side of the Coin ...... 6 II. MATERIALS AND METHODS ...... 11-26 III. PROGRESSIVE DISMANTLING OF THE HOST CYTOSKELETON FACILITATES APICOMPLEXAN PARASITE EXIT: TARGETING HOST PROTEASE ACTIVITY FOR DEATH BY ENTRAPMENT ...... 27-45 Tables 1-2 ...... 35-36 Figures 1-6 ...... 37-45 IV. HOST GPCR SIGNALING NECESSARY FOR PARASITE-MEDIATED CYTOLYSIS: THWARTING HOST SIGNALING TO STOP THEM IN THEIR TRACKS ...... 46-93 Tables 3-5 ...... 59-67 Figures 7-18 ...... 68-96 V. HUMAN PLATELET FACTOR 4 AND SYNTHETIC MIMICS: HARNESSING HOST DEFENSE MECHANISMS FOR PARASITE DEATH BY ULCER ... 97-119 Tables 6-8 ...... 106-108 Figures 19-24 ...... 109-119 VI. CONCLUSIONS AND FUTURE DIRECTIONS ...... 120-128 VII. BIBLIOGRAPHY ...... 129-150

LIST OF TABLES

Table 1. Proteins that are absent/reduced in infected erythrocyte and/or post-rupture vesicle membranes

Table 2. Proteases identified from infected erythrocyte and post-rupture vesicle membrane samples

Table 3. siRNA screen in U2OS cells Identifies Mediators of Parasite-Induced Cytolysis

Table 4. Multi-isoform families and components in initial siRNA screen.

Table 5. Isoform pair siRNA-mediated knockdowns.

Table 6. IC50 values determined in 3D7 (CQ-sensitive) and Dd2 (CQ-resistant) P. falciparum for 74 hits from smHDP screen.

Table 7. smHDPs show potency across several chloroquine-sensitive and chloroquine-resistant P. falciparum strains, with little cytotoxicity against mammalian cells.

Table 8. Lead smHDPs show potency across several chloroquine-sensitive and chloroquine- resistant P. falciparum field isolates.

LIST OF ILLUSTRATIONS

Figure 1. Fractionation strategy for comparing uninfected erythrocyte, infected erythrocyte and post-rupture vesicle membrane proteomes.

Figure 2. Comparative proteomic analysis of the erythrocyte membrane proteome during parasite egress.

Figure 3. Loss of adducin from the erythrocyte cytoskeleton prior to parasite escape is calpain- independent.

Figure 4. Proteomic profiling shows extensive cytoskeletal protein proteolysis during parasite egress.

Figure 5. Validation of calpain-dependent cytoskeletal processing during parasite rupture.

Figure 6. New model for parasite driven modifications to the host cytoskeleton during parasite egress.

Figure 7. siRNA screen identifies novel signaling components required for T. gondii-mediated cytolysis.

Figure 8. Design of siRNA screen to identify host genes required for T. gondii-mediated cytolysis.

Figure 9. Conserved function of host proteins in P. falciparum-mediated cytolysis.

Figure 10. Depletion of PLCβ1 and calmodulin-1 from erythrocytes is sufficient to block P. falciparum-mediated cytolysis.

Figure 11. Host PKC is activated late in the parasite intracellular cycle and abrogates host adducin cytoskeletal association.

Figure 12. Host Gαq and PLCβ1/γ1 are required upstream for host PKCα/β function in adducin loss prior to parasite-mediated cytolysis.

Figure 13. Host signaling cascade enhances calpain-mediated cytoskeletal proteolysis just prior to parasite exit.

Figure 14. Host [Ca2+] increases throughout the last third of the intracellular cycle, in a TRPC6- dependent manner.

2+ Figure 15. PKC activity, as mediated by Gαq and PLC, is required for Ca influx at the end of the parasite life cycle.

Figure 16. Mammalian PKC inhibitors show antiparasitic activity in vivo.

Figure 17. Parasite TCA cycle intermediates initiate the host cytolytic network.

Figure 18. Model of host GPCR-mediated cytolysis.

Figure 19. hPF4 acts as a HDP against P. falciparum via lysis of the parasite DV.

Figure 20. hPF4 causes dose-dependent DV lysis limited by protamine sulfate, and reduces parasitemia in a mouse malaria model.

Figure 21. Screen of smHDPs reveals potent inhibitors of P. falciparum growth.

Figure 22. smHDP leads kill P. falciparum via parasite DV lysis and decrease parasitemia in a murine malaria model.

Figure 23. smHDPs require a facial charge density to lyse the parasite DV.

Figure 24. Model of PF4 and smHDP uptake.

vii

INTRODUCTION

Apicomplexan Parasite Biology

Intracellular parasites of the phylum Apicomplexa cause significant morbidity and mortality worldwide. Toxoplasma gondii is a highly successful protozoan parasite, chronically infecting an estimated 30% of the world’s population (Weiss and Dubey,

2009). Including many water-borne protozoa that can threaten human health via accidental or intentional contamination of public water supplies, such as the class B biodefense agents Toxoplasma gondii, Cryptosporidium hominis, and Cryptosporidium parvum, this phylum is also a home to the malaria parasite Plasmodium falciparum, which continues to be a devastating disease and a global health burden responsible for

>500 million clinical cases of malaria that result in >1 million deaths each year (Gething et al., 2011). No completely satisfactory therapies exist for treating infection by these parasites: hypersensitivity to sulfonamides and lincosamides limits their utility against T. gondii, particularly in immunocompromised patients, and there is no effective chemotherapy for acute cryptosporidiosis. While vector control may reduce malaria parasite prevalence, chemotherapy remains the principal means of controlling disease.

Unfortunately, the emergence and spread of drug-resistant P. falciparum, the parasite that causes the most deaths, has rendered most of the traditional antimalarials (e.g. chloroquine) clinically ineffective, necessitating the discovery of novel antimalarials.

Ideally, novel treatments will minimize resistance, though most approaches to develop therapeutics generate resistant parasites in the lab and field.

Most Apicomplexan parasites are obligate intracellular pathogens that traverse a complex life cycle, including both sexual and asexual stages. The chronic asexual phase

1 includes a virulent lytic cycle responsible for parasite-mediated pathogenesis. During this lytic cycle, parasites (Plasmodium merozoites, T. gondii tachyzoites) divide inside a parasite-modified vacuole following host cell invasion (the parasitophorous vacuole or

PV). To do this they must dramatically remodel their host cell to facilitate invasion, intracellular development, avoid host defenses, and finally to exit from their host cell to renew the lytic cycle. Recently, there have been several studies investigating parasite proteins that interact with host proteins during the lytic cycle, but there is far less known about host proteins necessary for parasite growth.

Host-Pathogen Interactions Necessary for Survival: Getting In and Getting Out

The asexual phase of the intracellular cycle is comprised of a lytic cycle in which parasites invade a host cell and then establish an intracellular niche within a particular host cell type (Toxoplasma gondii infects any nucleated animal cell, Cryptosporidium parvum specifically infects enterocytes while Plasmodium falciparum infects erythrocytes). To initiate the intracellular cycle, parasites first attach to the host cell via low affinity interaction and actively reorient so that the end with apical organelles is pointed inwards towards the host cell. Parasite ligand interactions with host receptors cause irreversible attachment, which engages the parasite actin/myosin motor system to pull the host membrane around the parasite. Completion of invasion involves the sealing of the membrane to create a PV within which the parasite divides and completes its entire intracellular cycle. After replication is complete, the daughter parasites must exit from this PV and also the host cell in order to invade uninfected cells and continue the infection anew (Glushakova et al., 2005; Salmon et al., 2001). Many parasite-derived proteins, including proteases (Brossier et al., 2005; Buguliskis et al., 2010; Starnes et al.,

2009), kinases (Carruthers et al., 1999; Straub et al., 2011; Sugi et al., 2010) and

2 surface ligands (Brossier et al., 2003; Carruthers et al., 1999; Carruthers et al., 2000) have been implicated as necessary for invasion. Knockdown or inhibition of many of these proteins has been shown to prevent parasite invasion and cause parasite death.

Though parasite proteins required for invasion have been studied in detail, very little is understood on the host side, aside from a recent finding that invasion of T. gondii tachyzoites is dependent on host actin polymerization (Gonzalez et al., 2009), as mediated by interaction with a parasite-derived secreted actin-binding protein (toxofilin), which facilitates host cell actin filament disassembly and turnover (Delorme-Walker et al., 2012). Thus, invasion has proven to be a critical phase for therapeutic intervention, as parasites unable to invade are not only unable to replicate, these stranded parasites have been shown to die within hours of failed invasion (Buguliskis et al., 2010; Larson et al., 2009; Lovett and Sibley, 2003; Poupel and Tardieux, 1999; Sibley and Andrews,

2000; Teo et al., 2007).

Once a parasite successfully invades a host cell, it initiates major changes in its new environment. This is best illustrated by the extreme alterations P. falciparum parasites make to their new, and transient, home in a host erythrocyte. P. falciparum infection reduces deformability (Maier et al., 2008) and changes the adhesive properties of the host erythrocyte, due to the appearance of knob-like protrusions on the surface (Crabb et al., 1997; Sherman et al., 1992; Smith et al., 1992) and specialized secretion of parasite proteins into the host cell (Hiller et al., 2004; Marti et al., 2005). Yet perhaps the most profound change to the erythrocyte is its complete collapse at the end of the parasite erythrocytic cycle, as the newly replicated merozoites are released (Glushakova et al., 2005). For many years, parasite proteases have been thought to be important for rupture of malaria parasites from host cells (Hadley et al., 1983): from initial inhibitor-

3 based data, a two-step model of parasite rupture was proposed, requiring the activity of a cysteine and possibly a serine protease, as the general cysteine protease inhibitor E64 and the cysteine/serine protease inhibitors leupeptin and chymostatin inhibited PV and erythrocyte plasma membrane rupture, respectively (Glushakova et al., 2009; Salmon et al., 2001; Wickham et al., 2003a). More recently, several reports have implicated the involvement of three proteases in the rupture process: the parasite-derived proteases

PfDPAP3 (Arastu-Kapur et al., 2008) and PfSub1 (Yeoh et al., 2007), and the human protease calpain-1 (Chandramohanadas et al., 2009). In addition to proteolytic enzymes, it has also been established that osmotic forces (Glushakova et al., 2010), changes in parasite calcium flux (Nagamune et al., 2008), pore-forming proteins (Kafsack et al.,

2009), and a kinase (Dvorin et al., 2010) are important for efficient egress of apicomplexan family parasites.

Although these studies have provided details on a variety of parasite proteins that act or are proteolytically processed during egress (Bowyer et al., 2010), we hypothesized that the parasite likely modifies host erythrocyte proteins and specifically needs to destabilize the host cytoskeletal-membrane network to efficiently exit from the host cell,.

Thus, we were interested in a more comprehensive understanding of changes to the infected erythrocyte membrane and cytoskeleton proteomes as the parasites prepare the host cell for rupture and merozoite exit.

Egress of both P. falciparum (Wickham et al., 2003a) and the related apicomplexan,

T. gondii (Black et al., 2000; Lavine et al., 2007), has been shown to be a Ca2+- dependent process, inducing secretion of parasite-derived pore-forming proteins and activation of host-derived proteases. Ca2+ dynamics during egress have been best described within T. gondii: (i) It is well established that T. gondii egress can be induced via treatment with the calcium ionophore A23187 (Endo et al., 1982). (ii) T. gondii egress 4 has been shown to be controlled by a plant-like pathway for cyclic adenosine diphosphate ribose (cADPR) production through accumulation of abscisic acid, which results in elevated Ca2+ levels that stimulate egress (Chini et al., 2005; Nagamune et al.,

2008), and (iii) Ca2+-induced micronemal secretion of TgPLP-1, a perforin-like protein, has been shown to aid in lysis of the PV membrane during T. gondii egress (Kafsack et al., 2009). We have recently discovered that apicomplexan parasites co-opt the Ca2+- regulated host cysteine protease calpain to allow for egress from the host cell

(Chandramohanadas et al., 2009). In both P. falciparum and T. gondii, calpain was shown to be necessary for egress, as it is specifically activated just prior to parasite egress and is involved in the cleavage of several cytoskeletal proteins including α/β- spectrin and ankyrin. Pharmacological inhibition or depletion of calpain from host erythrocytes led to a robust block in parasite egress, thus limiting life cycle progression.

Recent work on T. gondii has implicated a perforin-like protein (TgPLP-1) as critical for egress (Kafsack et al., 2009). TgPLP-1 is a pore-forming protein similar to that used by cytotoxic T lymphocytes (CTL) for target cell killing (Voskoboinik et al., 2006). Perforin secretion by CTLs is thought to allow Ca2+ and granzyme entry, leading to target cell death (Chowdhury and Lieberman, 2008), which may link this parasite protein to host

Ca2+ signaling, as Ca2+ within the parasite may traverse this pore and mobilize into the host cell to activate a Ca2+ signaling pathway. TgPLP-1 KO parasites are crippled in vitro, and unable to establish infection in a mouse model, indicating the importance of this protein and the process of egress in the parasite survival. Deposition of this protein is necessary for efficient egress from host cells, as TgPLP-1 KO parasites fail to egress normally from the PV and host cell membranes. Strikingly, co-infection of host cells with both WT parasites and TgPLP-1 KO parasites abrogates the egress defect in the KO

5 parasite, thus implicating the likelihood of a parasite-induced freely diffusible second messenger in the host cell, which activates a common host-derived egress pathway allowing both parasites to exit.

Platelets, Malaria, and Host Defense Peptides (HDPs): the Other Side of the Coin.

In addition to interactions with the host that facilitate parasite survival, hosts have developed numerous defenses against these invading pathogens that have spanned evolutionary time, efficiently overcoming the barrage of potential invaders. Recent work has shown that platelets can bind infected erythrocytes and kill the intracellular malaria parasite (McMorran et al., 2009). Platelets have been long known to interact cooperatively with erythrocytes in thrombogenesis, both generally and in the specific case of sickle cell disease (Rocca and FitzGerald, 1997), though previous studies have suggested that thrombocytopenia is a poor prognostic marker in malaria (Moerman et al., 2003) and significantly associated with cerebral malaria (Cox and McConkey, 2010;

Wassmer et al., 2008).

Based on the knowledge surrounding possible interactions between platelets and P. falciparum and prior data that platelets contain HDP-like proteins (Tang et al., 2002), we hypothesized that perhaps innate immune proteins such as HDP-containing proteins that are found in either platelets could be natural antiparasitics and were likely optimized against these pathogens following years of concomitant evolution. Antiparasitic activities have been reported for a number of HDPs and are thought to kill protozoa, though the physiological significance of the reported interactions is questionable (Mangoni et al.,

2005; Vizioli and Salzet, 2002). The HDPs that have previously been studied, dermaseptins, are not present in blood or cells that circulate in the blood and kill intraerythrocytic parasites by disrupting the erythrocyte plasma membrane of infected

6 erythrocytes rather than directly killing parasites (Feder et al., 2000; Krugliak et al.,

2000).

HDPs play a central role in the innate immune system (Finlay and Hancock,

2004; Hancock and Lehrer, 1998; Tossi et al., 2000; Zasloff, 2002). There are two types of HDPs comprising ribosomally and nonribosomally synthesized peptides. Over 700

HDPs have been identified, varying in length (12 - 80 residues), sequence, and three- dimensional structure. The two major classes of HDPs are α-helical (magainin and cecropin) and disulfide-rich β-sheets (bactenecin and defensin), although other tertiary structures also exist. HDPs are particularly important in insects and plants, which lack an adaptive immune response based on T-cells and antibodies. Further, HDPs display very broad-spectrum action against bacteria, fungi, protozoa, and even viruses, which has promoted their use as new leads for developing antibiotics (Zasloff, 2002).

HDPs share common features including highly amphiphilic topologies in which the hydrophilic and hydrophobic side chains segregate onto distinctly opposing regions or faces of the overall folded conformation. Numerous studies with linear and cyclic peptides have strongly supported the hypothesis that the physicochemical properties of

HDPs, rather than any specific sequence or structure, are responsible for their activities.

It is generally believed that this amphiphilic topology is essential for insertion into and disruption of the cytoplasmic membrane leading to patogen death (Zasloff, 2002).

However, it is important to remember that while these compounds primarily target membranes, specific HDPs can interact with other components of the innate immune system (reviewed in (Finlay and Hancock, 2004)) and many HDPs appear to act on additional protein and nucleic acid targets (del Castillo et al., 2001; Kragol et al., 2001;

Park et al., 1998).

The cytotoxic activity and basis of specificity of cationic and amphiphilic peptides has been best studied in bacteria. This specificity for bacteria over mammalian cells is most likely related to fundamental differences between the two membrane types: bacteria have a large proportion of negatively charged phospholipid headgroups on their surface while the outer leaflet of animal cells is composed mainly of neutral lipids

(Zasloff, 2002). Also, the presence of cholesterol in the animal cell membrane appears to reduce the HDP activity. The bactericidal activity of these peptides is very rapid, occurring within minutes after exposure of bacteria to lethal doses of peptide. Several mechanisms have been proposed for the process of cell killing (Christensen et al., 1988;

Ludtke et al., 1995; Matsuzaki et al., 1995, 1996; Pouny et al., 1992). It appears that

HDPs bind to the membrane surface in a non-cooperative fashion and then aggregate once a threshold concentration is reached causing membrane permeabilization.

Membrane permeabilization is associated with a variety of other events including membrane depolarization, leakage of cellular metabolites, loss of compositional integrity due to enhanced membrane flip-flopping and translocation of peptides to the cytoplasm where additional targets, such as RNA, DNA, and various enzymes, can be encountered

(Scott et al., 2008). It is unclear which of these events are primarily responsible for bacterial killing and multiple hit models have been proposed with the exact mechanism being dependent on the peptide and bacterium in question (Patrzykat et al., 2002).

Nevertheless, HDPs have remained an effective weapon against bacterial infection over evolutionary time indicating that their mechanism of action thwarts bacterial responses, which normally lead to resistance against toxic substances. This premise is supported by data showing that no appreciable resistance to the action of the HDPs occurs after multiple serial passages of bacteria in the presence of sub-lethal concentrations of the peptides (Ge et al., 1999; Mosca et al., 2000; Tew et al., 2002). 8

Mature Platelet Factor 4/CXCL4, a member of the CXC chemokine family, is a 70 amino acid protein with an N-terminal cytokine domain, a central domain responsible for tetramerization, and a C-terminal domain that binds heparin and also can act as an HDP

(Yeaman et al., 2007). PF4 was the first characterized chemokine (Brandt et al., 2000), though unlike other chemokines, PF4 does not have strong chemotactic properties and does not interact with common chemokine receptors except human CXCR3B, an alternatively-spliced CXCR3 (Lasagni et al., 2003). PF4 is a cationic tetrameric molecule with high affinity for heparin and other large, negatively-charged molecules (Loscalzo et al., 1985). Upon platelet activation, PF4 is released from platelet alpha-granules in high local concentrations, up to 100 µM proximal to a clot (Lambert et al., 2007b). It is important to note that PF4 has been implicated in both inflammation (Scheuerer et al.,

2000; Srivastava et al., 2008) and thrombosis (Eslin et al., 2004b). The Poncz lab has shown that PF4 released from activated platelets forms complexes with endogenous

GAGs on the surface of platelets (Rauova et al., 2006) and monocytes (Rauova et al.,

2006) and likely on other hematopoietic and vascular cells. Mice that are knockout for the cxcl4 gene (Eslin et al., 2004b) (deficient in PF4 (PF4 KO)) or overexpress hPF4

(hPF4+) have proven to be important tools in understanding in vivo PF4 biology during thrombosis and megakaryopoiesis (Koenen et al., 2009; Lambert et al., 2007a).

Recently, Srivastava et al. have shown that PF4 mediates increased inflammation in a mouse model of experimental cerebral malaria (Srivastava et al.,

2008), which they posit may occur through the chemokine domain of PF4. It is unclear if these observations are clinically relevant to cerebral malaria in humans, given that this murine model (P. berghei ANKA) is complicated by severe inflammation in the brain and

CNS, unlike human cerebral malaria in which little, if any, inflammation is seen

(Bruzzone et al., 2009; Haldar et al., 2007; White et al., 2009). As human cerebral 9 malaria is a consequence of intravascular sequestration of infected erythrocytes in the brain vessels and capillaries, rather than increased inflammation, it is unlikely that PF4 mediates increased inflammation during this disease process. Murine models of cerebral malaria are further complicated by the marked accumulation of leukocytes and platelets in the brain venules and capillaries, a phenomenon not seen in human pathology (White et al., 2009). Given our preliminary in vitro and in vivo data implicating PF4 in

Plasmodium killing, utilizing P. falciparum in culture and a murine model that mimics systemic anemia during severe infections (P. yoelli), we hypothesize that in fact the HDP domain of PF4 serves as a natural human antimalarial and may offer insight into development of novel therapeutics with little propensity for generation of resistance.

II. MATERIALS AND METHODS

Reagents. Monoclonal antibodies against calpain-1 and Band 3 were purchased from Sigma.

Spectrin antibodies (α, β and β C-terminal domain) were donated by Dr. David Speicher (Wistar

Institute, Philadelphia), Band 4.9 antibody was purchased from BD Biosciences and all the other antibodies (α-Adducin, Ankyrin-1, Flotillin-1, Stomatin, Actin, and CD47) were purchased from

Santa Cruz Biotechnology. Calpastatin was purchased from EMD Biosciences. Recombinant human calpain-1 was purchased from Calbiochem. DCG04 was synthesized as reported previously (Greenbaum et al 2000). Streptavidin-HRP was purchased from Vector Labs, and

Fura-2-Dextran and SYTOX Green from Invitrogen. Purified human calpain-1 was purchased from Sigma, and calpastatin domain I from EMD Chemicals. Commercially available antibodies were obtained from the following sources: anti-OXGR1, anti-GPR91, and anti-TRPC6 from

Abcam; anti-ankyrin-1, anti-GAPDH, anti-calmodulin 1, anti-calmodulin 2, anti-CAMKI, anti-

CAMKII, anti-PKAcat, anti-PLCb, anti-PLCg from Santa Cruz; anti-PKCa and anti-PKCb from Cell

Signaling. Anti-α- and β-spectrin antibodies were kindly provided by D. Speicher (Wistar Institute,

USA).

Growth and maintenance of P. falciparum cultures. Strain 3D7 Plasmodium parasites were cultured in human erythrocytes (4% hematocrit) under standard conditions (37°C,

5%O2/5%CO2/90%N2) in RPMI buffer supplemented with HEPES and hypoxanthine. Replication was tightly synchronized via serial treatments with D-sorbitol to lyse trophozoite- and schizont- infected erythrocytes. The remaining ring-stage parasites were cultivated to yield schizonts (~42 hr post-infection), which were selectively purified using a Miltenyi Biotec magnetic separator.

Cultures were staged by microscopy of Giemsa-stained blood smears and flow cytometry using

SYTOX Green (Life Technologies) as a marker of DNA content. Cultures were fixed with 4% paraformaldehyde/0.0016% gluteraldehyde for 1 hour and permeabilized with 0.1% Triton X-100 for 10 minutes at room temperature prior to incubation with 5 µM SYTOX Green for 10 minutes

11 and analysis on an Accuri Flow Cytometer.

Growth and maintenance of T. gondii cultures. RH strain T. gondii tachyzoites, and parasite lines constitutively expressing P30-GFP or P30-RFP were cultivated in confluent monolayers of human foreskin fibroblasts, human U2OS cells, or human A549 cells. Cells were grown to confluence in Dulbecco’s modified Eagle’s medium containing 10% fetal bovine serum and replaced upon infection with minimal essential medium containing 10% dialyzed fetal bovine serum.

Preparation of erythrocyte membrane/cytoskeletal fractions. To prepare the infected erythrocyte membrane, schizont-stage infected erythrocytes (∼35-40 hpi) were purified using a magnetic column (Miltenyi Biotec). The parasites were then treated with 0.02% saponin in PBS for 10 min at 4°C and centrifuged at 1,500xg. The supernatant was centrifuged again at 4,000xg.

This supernatant containing infected erythrocyte and parasitophorous vacuole (PV) contents was then ultracentrifuged (200,000xg for 2 hour at 4°C) to collect membranes. The membrane/cytoskeletal pellet was washed three times with PBS and subjected to a delipidation procedure by ethanol extraction (Pasini et al., 2006) for 90 min prior to freezing with protease inhibitors (Roche) at -800C. Finally, uninfected erythrocyte membranes were prepared as mentioned previously (Pasini et al., 2006). Erythrocytes were washed twice with phosphate buffered saline (PBS) and subjected to hypotonic lysis in 5 mM lysis buffer (5 mM K2HPO4, pH

8.0) containing protease inhibitors for 10 min in ice. Ghost membranes were collected by centrifugation of the lysate (200,000xg for 2 hour at 4°C). The pellet fraction containing the erythrocyte membrane was washed until pellet appeared as yellow/white ghosts. After delipidation, the membrane samples were frozen until used.

Purification and microscopic analysis of the erythrocyte-derived vesicles formed during rupture. Schizont-stage parasites ~40 hours post invasion (hpi) were purified from synchronous

12 cultures using a SuperMacs magnet (Miltenyi Biotech) and resuspended in culture medium. Once the majority of schizonts ruptured, as determined by Giemsa smears, the culture media was treated with protease inhibitor cocktail and further collected by centrifugation at 1,500xg for 10 min to remove residual uninfected and infected erythrocytes. This was followed by centrifugation at 5000xg for 30 min to remove extracellular merozoites. Erythrocyte membrane material, including small vesicles and loose membranes, resulting from rupture of parasites from erythrocytes was then purified from the residual media by ultracentrifugation (L8-70M ultracentrifuge, Beckmann) at 200,000xg for 2 hour at 4°C. The resulting erythrocyte membrane material was then washed three times with PBS. The post-rupture erythrocyte vesicles isolated were stained with FM4-64 (Invitrogen) and DAPI for DNA. Samples were analyzed using a fluorescent microscope (Carl Zeiss Axioskop). Composite images were produced using

Photoshop 6.0.1.

Protein separation for mass spectrometry (MS). Aliquots from uninfected, infected erythrocytes and post-ruptured vesicle membranes (∼100 µg from each sample) were solubilized in Laemmli buffer under reducing conditions. After resolving on a 4-16% pre-cast gradient gel

(BioRad), polypeptides were stained with colloidal Coomassie. For mass spectrometric purposes, gel slices of 0.5 cm were excised from the entire length of the gel for each sample (total of 34 slices). Gel slices were de-stained with 20 mM ammonium bicarbonate and dried in 100 % acetonitrile. Tryptic peptides from each gel slice were dried on a vacuum evaporator for mass spectrometry.

Mass Spectrometry. These studies were completed by Angela Wehr, Ph.D. Tryptic peptides were resuspended in acetonitrile/water. Peptides were separated by reverse phase using an

Agilent 300 Extend-C18 column (3.5mm, 2.1 x 150mm). After a 6 min isocratic hold at 100% mobile phase A linear gradient was run over the next 33 minutes increasing the percent B (0.1% formic acid, 95% acetonitrile) to 60% all at a flow rate of 150 mL/min. Peptides were eluted online to a ThermoFinnigan LTQ mass spectrometer (ThermoFisher, San Jose, CA) and were ionized

13 using ESI in the positive mode. Nitrogen was used as the sheath gas and collision induced disassociation (CID) experiments employed helium. Data dependent scanning was used to identify the top five most abundant ions over the range m/z 400-1500. Each of the five most abundant ions was then subjected to product ion scanning. (A mass tolerance of 2.0 Da for precursor ions and 1.0 Da for fragment ions was permitted). We used the extract_msn.exe tool from BioWorks version 2.x for generating peak lists from individual spectra.

Processing of Mass Spectrometry Data. These studies were completed by Angel Pizzarro,

Ph.D. at the Institute for Translational Medicine and Therapeutics at the University of

Pennsylvania. The raw MS/MS data were searched through SEQUEST (TurboSEQUEST - PVM

RefSeq database (NCBI version updated 11/19/07) or the Plasmodium database (plasmodb.org).

The databases were indexed using strict trypsin cleavage rules (K and R) with two maximum internal cleavage sites. Carbamidomethylation of cysteine residues (fixed modification) and oxidation of methionine (variable modification) were pre-set for the database searches. The

SEQUEST output files were analyzed and validated by PeptideProphet (Version 3.0) as an automated method to assign peptides to MS/MS spectra. Post-analysis of the data was performed using the Trans Proteomic Pipeline (TPP) with a peptide identification probability cutoff of 0.05, which for all data sets corresponded to a FDR < 3%, and was < 2% for most data sets.

Data was further filtered and proteins with at least three peptides identified by SEQUEST were considered as positive ‘hits’. Multiple biological experiments were carried out to reproduce and confirm the proteomic results. GeneSpring Version 11.5 (Agilent) was used for generating heat map from the information obtained from MS analysis to analyze the relative distribution of proteins in gel slices among the three proteomes.

Immunodepletion and loading of erythrocytes. Erythrocyte ghosts were prepared by hypotonic

lysis as previously described utilizing 5 mM K2HPO4 at <4°C for 15 minutes (Chandramohanadas, et al 2009). To immunodeplete target proteins, target antibodies were pre-conjugated to Protein

G-Sepharose (Upstate/Millipore) following a titration to achieve maximal immunodepletion (1-10

µg antibody per 106 erythrocytes). Conjugates were incubated with ghosts for 3 hours on ice with gentle mixing and repeated with fresh antibody-sepharose conjugates up to 3 times. The slurry was separated by centrifugation and followed by another round of immunodepletion for cells immunodepleted of multiple proteins. Ghosts were resealed by gradual addition of 5X resealing buffer (475 mM KOAc, 25 mM Na2HPO4, 25 mM MgCl2, 237.5 mM KCl, pH 7.5) over 1 hr at

37°C. Parallel studies were carried out using anti-PKAcat, and mock treated erythrocytes were incubated with Sepharose G beads without antibodies.

Analysis of host protein function during parasite exit from immunodepleted erythrocytes.

Resealed erythrocytes were prepared as described above. Schizont stages were isolated from 50 ml parasite culture by magnet purification ~40 hr after sorbitol synchronization, and added to mock-treated, PKCa/b-depleted, PLCb1-depleted, calmodulin-1-depleted, Gaq-depleted, or calpain-1-depleted erythrocytes to a final hematocrit of 4%. Parasite progress through the intraerythrocytic cycle was monitored by Giemsa smear. Schizont-stage parasites were followed to assess egress and the establishment of rings in newly-infected erythrocytes from 45-60 hpi.

Flow cytometry was used for quantitative evaluation of P. falciparum development. 106 events were collected per sample using an Accuri flow cytometer and analyzed using Accuri software, gating to exclude debris defined by scatter characteristics, and uninfected erythrocytes based on low fluorescence; rings and trophozoites were distinguished from schizonts based on DNA content. Data are presented as mean ± SEM; n=4.

Mechanical removal of P. falciparum from erythrocytes. Needle-shearing experiments were completed as previously described (Dvorin, et al 2010) with modifications. Parasites trapped within erythrocytes immunodepleted of calpain-1, PKCa/PKCb, PLCb1, or CaM-1, or negative control Mock-immunodepleted or PKAa/bcat-immunodepleted erythrocytes, were plated in quadruplicate. At the indicated timepoint, cultures were sheared by 20 strokes through a 28.5

15 gauge needle. Cultures were incubated for an additional 12 h and ring stage parasitemia determined by Giemsa smear microscopy and flow cytometry of DNA content. Data are presented as mean ± SEM; n=4.

Monitoring calpain activation and membrane binding. Immunodepleted cells were prepared as above and challenged with synchronous, magnet-purified schizont stage parasites. Following

48-hour incubation, infected cells were incubated with DCG04 (5 µM) and membrane fractions were prepared from each sample by ultracentrifugation (2 h at 200,000xg). Equal amounts of solubilized protein from each sample was separated by SDS-PAGE, transferred to PVDF membrane, and probed for biotin using streptavidin-HRP, revealing DCG04 labeling of active calpain. As a loading control, the same western blots were probed with actin antibodies (Santa

Cruz).

Erythrocyte membrane proteolysis in vitro by calpain-1. Aliquots of 10 µl (1µg/ µl) erythrocyte membrane preparations were treated with 100 ng of purified calpain-1 in reaction buffer (20 mM

HEPES, 10 mM DTT & 5 mM CaCl2) for 30 min at 37°C. Negative controls were included by pre- incubating erythrocyte membranes with calpastatin or EGTA prior to the addition of activated calpain. Samples were solubilized in Laemmli buffer under reducing conditions and resolved via

SDS-PAGE and visualized by colloidal Coomassie. A fraction from each reaction was also resolved on SDS-PAGE, transferred to PVDF membrane and probed for α- and β-Spectrins,

Ankyrin-1, Stomatin, and Flotillin using commercially available antibodies.

Western Blot Analyses. Protein samples were solubilized in Laemmli buffer, separated by SDS-

PAGE and transferred to PVDF membranes. Membranes were probed using available antibodies and visualized with corresponding secondary antibodies conjugated to HRP (Pierce). Hypotonic lysis and resealing of erythrocytes with calpastatin was done as previously published [15].

Proteomic Protein Profiling. These studies were completed in collaboration with Rajesh

Chandramohanadas, Ph.D. Protein degradation was assessed by creating “chromatograms”

16 where the number of peptides identified for each protein was plotted against the corresponding gel slice where it was identified (as an approximation of molecular weight) from uninfected, infected and post-rupture erythrocyte membrane proteomes. Majority of proteins peaked in gel slices that roughly corresponded to the nominal molecular weight among the three proteomes.

Appearance of “peptide peaks” in lower molecular weight areas of the gel in the post-rupture proteome and a corresponding absence or lessening of this peak in both uninfected and infected samples suggested protein degradation in the former sample. To further evaluate protein processing, peptides identified from proteolytic fragments were aligned manually against peptides identified from the full-length protein (identified at the correct molecular weight area of the gel), for several proteins that were found processed using chromatograms.

Immunofluorescence studies. Synchronous P. falciparum-infected erythrocytes or P30-

GFP/P30-RFP T. gondii-infected U2OS cells were fixed for 1 hour at RT with 4% paraformaldehyde/0.008% gluteraldehyde, washed three times with PBS, permeabilized using

0.1% Triton X-100 for 10 minutes at 37°C, and blocked using 5% milk in PBS for 1 hour at RT.

Cells were incubated with primary antibodies to adducin-1 (Santa Cruz) in 5% milk in PBS overnight at 4°C, washed three times with PBS, and incubated with secondary antibodies preconjugated to Alexa-568, Alexa-Cy3, or Alexa-488 (Life Technologies) for two hours at RT.

Cells were washed three times prior to incubation with Hoechst DNA stain and imaging on a

Leica DMI6000B epifluorescent scope.

Monitoring T. gondii tachyzoite invasive capacity following block in host cell cytolysis.

Utilizing a T. gondii RH parasite line that expresses firefly luciferase, infected cells were mechanically disrupted via scraping and filtration to remove host cell material at specified time points following tachyzoite arrest within A549 host cells (beginning at 40 hpi) expressing

S716A/S726A mutant adducin. Parasites were then allowed to invade confluent monolayers of

HFF cells in 96-well plate format and assessed for viability 12 hours later utilizing the Luciferase

Reporter Assay Kit (Biovision) according to manufacturer’s instructions. Parasite viability was 17 assessed relative to that of tachyzoites that originally infected wild-type A549 cells prior to mechanical release. Data shown are means of at least 3 separate experiments +/- SEM.

siRNA Screen and shRNA stable knockdown generation. Primary siRNA oligos were purchased from Santa Cruz, oligos for follow up screens were purchased from Ambion. siRNAs were added at a final concentration of 20 nM in 96-well plates and titrated to achieve maximal knockdown by 72 hours post-transfection, along with 106 U2OS cells in culture medium lacking phenol red. Reverse transfection was carried out using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s instructions. Transfected host cells were inoculated with 105 P30-GFP- expressing T. gondii tachyzoites 24 hr post-transfection, and followed throughout the intracellular cycle to assess invasion, replication, and egress rates at 6, 24, 44, and 60 hpi. Data are presented in Figure 1B and C as mean number of unruptured vacuoles at 60 hpi versus 44 hpi ±

SEM; n=4. siRNA knockdown was confirmed via western blot of mirrored cultures utilizing validated antibodies from Santa Cruz. Stable knockdown cell lines were generated following transfection with huSH shRNA plasmids (Origene) encoding target shRNAs conjugated to soluble cytoplasmic RFP to affirm knockdown and selection with puromycin (1 µg/ml) for 2 passages, prior to isolation of clonal populations. Double knockdown cells were generated utilizing pGFP-B-

RS expression vectors (Origene) and selection with blasticidin. Following confirmation of knockdown via western blot, cells were cultured in the absence of selection pressure.

T. gondii invasion, replication, and egress in knockdown cultures. Intracellular parasite growth was measured by counting the number of parasites per parasitophorous vacuole at 6, 24 and 44 hr post-infection (prior to egress from the initial host cell). Confluent monolayers of ~5 x

105 stable knockdown host cells in 60 mm dishes were infected with 106 T. gondii parasites. At least 100 vacuoles were counted per time point. Data are presented as mean ± SEM; n=5.

Monitoring T. gondii egress upon treatment with TCA metabolites. P30-GFP T. gondii

18 tachyzoites were incubated in quadruplicate with confluent monolayers of HFF cells within glass- bottom 35 mm dishes and allowed to undergo 3 parasite divisions (8 tachyzoite per PV stage) prior to the addition of 50 µM lactate, aKG, or succinate, or a mixture of αKG and succinate, or the Ca2+ ionophore A23187 (10 µM) as a positive control of egress induction, to the culture media. Dishes were followed by fluorescent microscopy within an environmental chamber that mimics incubator conditions (37 degrees C, 5% CO2) until parasite egress.

P. yoelii 17XNL and P. berghei ANKA mouse studies. These experiments were conducted with the assistance of Satish Mishra, Ph.D. in the laboratory of Photini Sinnis, M.D. at the Malaria

Research Institute, Bloomberg School of Public Health, Johns Hopkins University.

Swiss Webster mice were infected with 2x105 P. yoelii 17XNL parasitized erythrocytes in

200 µL of RPMI medium via intravenous injection. Following a standard 4-day Peters suppression test protocol (Knight and Peters, 1980), the treatment regimen began on day 1 (the same day the mice were infected) and mice were dosed once per day intravenously on days 1-3 with vehicle

(20% DMA in saline [n=3] or 20% Kleptose HPB [n=2]); 10 mg/kg mPF4 (in saline [n=3], R&D systems); or 10 mg/kg C12 (in saline [n=4]). Parasitemias were determined on day 4 via Giemsa- stained blood smears.

C57BL/6 mice were infected with 2.5x104 P. berghei ANKA parasitized erythrocytes via intravenous injection. Mice were dosed once per day intravenously on days 1-4 with either 5 mg/kg PMX1207 (in 20% DMA in saline [n=5]); 20 mg/kg PMX207 (in 20% Kleptose HPB [n=5]); or untreated (Control, n=5). Parasitemias were determined on days 5 and 7 via Giemsa-stained blood smears.

T. gondii mouse studies. These studies were completed by Christopher D. Dupont in the laboratory of Christopher A. Hunter, Ph.D. at the School of Veterinary Medicine, University of

Pennsylvania. Age-matched female Swiss Webster mice were infected intraperitoneally with 103

RH parasites engineered to express the model antigen Ovalbumin and the fluorescent protein

Tomato (RH-OVA-Tom; John, et al. 2009). Treated mice were administered 10 mg/kg Go6976 daily beginning 1-hour post-infection. Five days post-infection, mice were euthanized, and parasite burden in the spleen and peritoneal cavity was determined by flow cytometry. Statistical significance was determined using a two-tailed Mann-Whitney Test. Data shown are compiled from three separate experiments. Experiments were performed at the University of Pennsylvania

(UPenn) School of Veterinary Medicine in accordance with the guidelines of the UPenn

Institutional Animal Care and Use Committees.

PECS were obtained by peritoneal lavage with 5 mls of PBS (GIBCO). Splenocytes were obtained by mashing spleens against a 40 µm cell strainer (Biologix) and resuspending in complete RPMI (RPMI 1640 (Mediatech), 10% fetal calf serum (GIBCO), 1X non-essential amino acids (GIBCO), 1X Penicillin Streptomycin (GIBCO), 1X beta-2-mercaptoethanol (GIBCO)). Red blood cells were lysed by resuspending in lysis buffer (0.846% NH4Cl (Sigma-Aldrich)). Cells were suspended in flow cytometry buffer (PBS, 0.2% bovine serum antigen (Sigma-Aldrich), 2mM

EDTA (Invitrogen)) and analyzed using a FACSCanto (BD) or LSRII (BD) flow cytometer. Data was analyzed using FlowJo 8.8.7 software (TreeStar) and Prism 4 software (GraphPad).

CaMKII studies. U2OS cells were fractionated using Triton X-100. The cytoskeletal fraction was incubated with 100 nM recombinant rat CAMKII (NEB) in 1X NEBuffer for Protein Kinases supplemented with 200 µM ATP, 1.2 µM calmodulin and 2 mM CaCl2 and incubated for 1 h at

37°C. 100 nM recombinant human calpain (Biovision) was added to the mixture and incubated for

30 min at 37°C prior to separation by SDS-PAGE, transfer to PVDF membrane, and probing for a-spectrin (primary antibody gift from David Speicher) and anti-rabbit secondary antibodies conjugated to HRP (Sigma).

Ca2+ studies. Erythrocytes were hypotonically lysed and loaded with 50 µM fura2-dextran prior to resealing and incubation with synchronous P. falciparum schizonts. Parasites infecting fura-2- dextran-loaded erythrocytes were followed by fluorescent microscopy and plate-based

20 fluorometry throughout the life cycle to assess host erythrocyte cytoplasmic [Ca2+] via the

2+ equation: [Ca ] = Kd Q (R-Rmin)/ Rmax-R), where R represents the fluorescence intensity ratio at

340 nm/380 nm excitation and emission at 510 nm. Data are presented as mean fluorescence ±

SEM; n=3. Representative images are shown.

U2OS cells were transduced using the Premo Cameleon Calcium Sensor (Life

Technologies) according to manufacturer’s instructions to facilitate cytoplasmic expression of

YC3.6 prior to synchronous T. gondii infection. Host [Ca2+] was measured throughout the parasite life cycle via FRET microscopy on a Leica DMI6000B and FRET-based fluorometry to assess

[Ca2+] as a function of CFP/YFP emission ratio. Data are presented as mean fluorescence ±

SEM; n=3. Representative images are shown.

Assessing background (digestive vacuole) FRET-based or Fura-2 fluorescence. In order to determine CKAR or Fura-2 signal that was attributable to uptake into the parasite digestive vacuole, sorbitol-synchronized 3D7-CKAR parasites or 3D7 parasites infecting erythrocytes loaded with Fura-2 dextran were selectively removed from their host cells using 0.1% saponin in

PBS at 3 hour increments throughout the intracellular cycle, washed, and plated in quadruplicate prior to assessment of plate-based fluorescence as described in the main experimental procedures. Representative images were also taken on a Leica DMI6000B epifluorescent microscopy. Data are presented as mean +/- SEM; n=4.

CKAR transfection and PKC activity measurement. CKAR plasmid (Addgene) was transfected into U2OS cells using Lipofectamine 2000 (Life Technologies) according to manufacturer’s instructions. 3D7-CKAR construct was generated via addition of a minimal export PEXEL motif N- terminal to CKAR (Marti, et al 2004; Hiller, et al 2004) following PCR amplification from the CKAR plasmid. This was achieved via amplification of the N-terminus of the exported protein KAHRP

(PF3D7_0202000) from cDNA of asynchronous 3D7 parasite cultures and recombination with the

CKAR motif into the P. falciparum expression vector p-HHVPatt using MultiSite Gateway (Marti et

21 al., 2004). Constructs were transfected into the P. falciparum 3D7 line and selected using 10 nM

WR99210 (an inhibitor of dihydrofolate reductase; Fidock and Wellems, 1997). 3D7-CKAR transgenic parasites and T. gondii-infected U2OS cells expressing CKAR were cultured under standard conditions and assessed for PKC activity via FRET-based fluorometry as a function of

YFP/CFP emission ratio and FRET imaging on a Leica DMI6000B. Data are presented as mean

± SEM; n=4. Representative images are shown.

Parasite IC50 determination. These studies were completed in collaboration with Melissa S.

Love at the University of Pennsylvania. Compounds were assayed for 72h, after which the cultures were fixed with a solution of 4% paraformaldehyde and 0.008% glutaraldehyde in PBS prior to permeabilization with 0.25% Triton X-100, and 5 µM SYTOX Green Nucleic Acid Stain and flow cytometry analysis (Accuri C6 Flow Cytometer with C-Sampler). IC50 curves were generated using GraphPad Prism.

Hemolysis. These studies were completed by Melissa S. Love at the University of Pennsylvania.

Following removal of culture supernatant before fixing and transferring to a clear-bottom plate.

Absorbance at 541 nm was measured and compared to a standard curve generated from Triton

X-100-lysed erythrocytes.

Platelet activation. Platelets were collected from C57BL/6 mice (WT), homozygous PF4 knockout mice (PF4 KO), and transgenic mice overexpressing human PF4 (hPF4+) (Eslin et al.,

2004a) and added to cultures either with or without pre-activation ex vivo with 5 µM AYP.

Parasitemia was assessed via flow cytometry after 24 hours treatment.

Parasite membrane potential assays. These studies were completed in collaboration with

Melissa S. Love at the University of Pennsylvania. Parasite-infected erythrocytes were incubated 22 at 37°C for 30 minutes with 1 µM rhodamine 123 (for parasite plasma membrane potential), 0.2

µM rhodamine 123 (for parasite mitochondrial potential) (del Pilar Crespo et al., 2008), or 10 nM

LysoTracker Red (for digestive vacuole potential). After pre-incubation, parasites were treated for

4 hours with the test compound, a 10 µM mixture of the ionophores monensin (Sigma) and nigericin (Calbiochem) as a positive control of membrane potential perturbation, or left untreated and then analyzed via fluorescence microscopy on a Leica DMI6000 B.

Fluorescence microscopy and quantification of digestive vacuole lysis. Erythrocytes infected with synchronous cultures of PMII-GFP expressing parasites were treated at 30 hours post invasion (hpi) and fixed at the indicated time points, stained with Hoechst and imaged by fluorescence microscopy. Cells were also analyzed on an Amnis ImageStream X high-resolution flow cytometer with gating for Hoechst-positive cells to isolate infected cells (Muskavitch et al.,

2008; Ortyn et al., 2007).

smHDP screen. These studies were completed in collaboration with Melissa S. Love at the

University of Pennsylvania. Synchronized parasites were treated with 500 nM of each smHDP for

72 hours. Cells were then fixed and assayed for parasitemia via flow cytometry. Compounds were deemed hits if they caused ≥80% parasite death, as compared to a 500 nM chloroquine-treated

3( + ) Z' = 1 ! p n positive control. The Z-factor was calculated using the equation p ! n where µ represents the means and σ represents the standard deviations of the positive (p) or negative (n) controls (Zhang et al., 1999). !

HDP screen and assay for hemolysis. These studies were completed by Melissa S. Love at the

University of Pennsylvania. P. falciparum parasites (3D7 strain) were cultured as described previously. Synchronized parasites were treated with 15 µM of 9 platelet-, neutrophil-, or

23 lymphocyte-derived proteins containing HDP-domains or properties: Regulated upon Activation,

Normal T-cell Expressed, and Secreted (RANTES, ProSpec); Platelet Factor-4 (PF4,

Haematologic Technologies Inc.); Fibrinopeptide-A (FP-A, AnaSpec); Fibrinopeptide-B (FP-B,

AnaSpec); Human Neutrophil Peptide-1 (HNP-1, AnaSpec); Human Neutrophil Peptide-2 (HNP-2,

AnaSpec); Cathelicidin (LL-37, AnaSpec); Lymphotactin (ProSpec); Lactoferrin (ProSpec). 15 µM melittin, the major active component of bee venom, was used as a positive control for HDP-killing of parasites with complete hemolysis. Parasite death was normalized to a 500 nM artesunate control. Parasites were treated for 72 hours, and then fixed and assayed for parasitemia via flow cytometry. Hemolysis was measured by removing 50 µL from each well (100 µL total in a 96-well plate) before fixing and transferring to a clear-bottom plate. Absorbance at 541 nm was measured and compared to a standard curve generated from Triton X-100-lysed erythrocytes (4% hematocrit serially diluted by 2).

Fluorescence microscopy of PMX496. Erythrocytes infected with synchronous cultures of wild type parasites (3D7) were treated at 30 hpi with 250 nM of the intrinsically fluorescent PMX496

(green: PMX496, blue: parasite nuclei) and followed at specified time points on a Leica DMI6000

Protamine sulfate studies. hPF4 immunofluorescence was assayed as described above with pretreatment of cultures with 5 mM protamine sulfate or mock (DMSO) for 30 min prior to hPF4 treatment.

Western blot analysis of PF4 accumulation. Synchronous cultures of P. falciparum-infected erythrocytes were treated with 10 µM PF4 then 0.1% saponin to separate erythrocyte fractions from parasite material. Samples were run by standard SDS-PAGE, transferred to Immobilon

Transfer PVDF membrane and incubated with 1:1000 primary antibody to human PF4 and

1:10000 anti-mouse secondary antibody. Input of total cell lysates was used as a loading control, using a primary antibody to flotillin and an anti-rabbit secondary antibody.

Transmission Electron Microscopy. Synchronous cultures of P. falciparum parasites were magnet-purified at ~25 hpi and allowed to sit for 6 hours prior to a 10-min exposure to 10 µM

PF4, 500 nM PMX207 or PMX1207. DMSO was used as a negative control. Samples were frozen in an Abra high-pressure freezer. This material was freeze substituted in 2% OsO4 in anhydrous acetone and embedded into Epon. Images were collected at 120KeV on a FEI Tecnai12 equipped with a Gatan 894 2k camera. Images were captured by Dewight R. Williams, Ph.D. at the Electron Microscopy Core at the University of Pennsylvania. Tilt series were taken +/- 70° in

3° increments at 6500x magnification. Tilt images were decimated by a factor of 2 and initially aligned by cross-correlation in FEI’s inspect3D. Tomograms were constructed from fiducially aligned stacks, orthogonal merged axes, and subsequent serial tomograms joined in the IMOD package “etomo”. Montage images taken with 10x10 sampling at 3200x magnification were aligned with blendmont.

Texas Red Dextran-loaded erythrocyte analysis of DV lysis. Erythrocytes were lysed and resealed as described previously in the presence of 10 µM Texas Red Dextran prior to challenge with synchronous magnet-purified cultures of P. falciparum-schizont stage parasites. Infected erythrocytes containing Texas Red Dextran were treated at 30 hpi with hPF4, C12, or smHDPs as previously described and imaged on a Leica DMI6000B epifluorescent microscope to assess lysis of the parasite DV, denoted by Texas Red Dextran dispersion throughout the parasite body.

Infected Texas Red Dexran-loaded cells were also subjected to gametocytogenesis as previously described (Fivelman et al., 2007) and treated with hPF4 or PMX1207 at stage 2 prior to fluorescent microscopy analysis of DV lysis.

Statistical analyses. Statistical significance of unruptured vacuole persistence, in vitro parasitemias, FRET-based signal for CKAR activity or YC3.6 Ca2+ studies, and Fura-2 signal with a one-way ANOVA with Dunnett’s post-hoc test. In vivo mouse parasitemias were analyzed with one-tailed paired Student’s t-tests and mouse survival was analyzed using the Mantel-Cox log- rank test.

III. Progressive dismantling of the host cytoskeleton facilitates

Apicomplexan parasite exit:

Targeting host protease activity for death by entrapment

We employed a proteomic approach to understand the molecular basis for the large changes in the infected erythrocyte cytoskeleton upon schizont maturation and egress.

Three proteomes were compared via mass spectrometry-based analysis: (i) the plasma membrane proteome from uninfected erythrocytes, (ii) the erythrocyte/PV membrane proteome from infected erythrocytes prior to parasite egress (~40 hpi, 8 hours prior to egress), and (iii) the erythrocyte/PV membrane proteome isolated from infected erythrocyte post-rupture vesicles. Although there certainly could be important cytosolic protein modifications during parasite egress, changes in the soluble proteome were largely obscured by hemoglobin, thus we specifically analyzed proteins in only the membrane fractions.

Proteomic analysis of changes to the host membrane proteome during parasite egress

We developed a differential fractionation strategy to isolate these three host membrane proteomes (Figure 1A). Uninfected erythrocyte membranes were isolated by hypotonic lysis followed by ultracentrifugation. To isolate extraparasitic membranes

(PV/erythrocyte) from infected erythrocytes, magnet-purified schizonts were separated from uninfected erythrocytes (Greenbaum et al., 2002), then treated with saponin, which at low concentrations (0.02%) selectively permeabilizes the erythrocyte/PV sparing the parasite plasma membrane (Chandramohanadas et al., 2009). The erythrocyte/PV membranes were then enriched by low speed centrifugation to remove whole parasite 27 cells and further isolated by ultracentrifugation as described elsewhere

(Chandramohanadas et al., 2009). To isolate post-rupture vesicle membranes from infected erythrocytes, highly synchronous cultures of late-stage infected erythrocytes were magnet-purified and allowed to naturally egress in culture media. Extracellular merozoites were then separated from culture media (containing post-rupture membrane vesicles) by low speed centrifugation (1,500xg). Post-rupture membrane vesicles were then isolated from the media by ultracentrifugation at 200,000xg for 2 hours. For comparison, we show FM4-64 stained images of uninfected erythrocytes; parasite culture media collected immediately after rupture that contains both intact uninfected erythrocytes and post-rupture vesicles (red), as well as released merozoites stained with

Hoechst (blue); and purified post-rupture vesicles purified from the media after parasite egress (red) (Figure 1B). We observed little parasite contamination in either the infected erythrocyte membrane preparation or the samples enriched for post-rupture vesicles. To confirm that we achieved purification of extraparasitic membrane material, we show the enrichment of the erythrocyte membrane marker Band 4.9 and the concomitant loss of

GFP from a parasite line expressing cytosolic GFP (Figure 1C).

To identify and compare proteins from these three proteomes we employed an SDS-

PAGE/MS approach akin to the geLC-MS/MS approach used by the Mann group

(Shevchenko et al., 2006), where samples were first separated by one-dimensional SDS

PAGE, then individual gel slices across the entire lane were excised and subjected to

LC/MS/MS analysis. The uninfected, infected erythrocyte and post-rupture vesicle membranes were first delipidated by incubating with sodium acetate/ethanol for 1 hour at room temperature, which enhanced the recovery of peptides in the subsequent LC/MS analysis (Pasini et al., 2006). The polypeptides were then separated by SDS-PAGE and

28 stained with colloidal Coomassie. Initially we observed that the uninfected erythrocyte and the infected erythrocyte membrane proteome migration pattern by Coomassie to be comparable, apart from a major reduction in spectrin band intensity in the infected erythrocyte membrane sample. However, the migration pattern for the post-rupture vesicle membrane proteome was markedly different (Figure 2A).

Thirty-four individual gel slices were excised throughout the SDS-PAGE lanes in order to identify proteins and assess their relative size for each of the three proteomes

(as graphically depicted in Figure 2A). Each excised gel band was digested with trypsin, peptides were analyzed by LC/MS/MS and peptides were matched to proteins using

SEQUEST. A total of 33,386 entries from the NCBI database were searched for protein identification. Proteins that had at least three peptides from a minimum of two independent MS analyses were considered as confident ‘hits’. Most datasets had a false discovery rate of < 2. Although the Coomassie stained gels only revealed a few major proteins, we identified approximately 250 proteins from each sample (Figure 4B), which is commensurate with prior proteomic studies of human erythrocytes (Pasini et al.,

2006); thus most proteins were present at a level below the level of detection by

Coomassie. Furthermore, proteins identified were well distributed throughout the SDS-

PAGE gel (Figure 2B). Spectral counting analysis was used to compare the ratios of total protein abundance, which revealed that only ~15% of the protein hits changed more than 3-fold (first two bars in histogram), while the relative abundance of the remaining

~85% of proteins was within 3-fold (0.3-1 and 1-3) amongst all three proteome samples.

This data indicates that although the Coomassie stained gels showed different patterns- primarily due to huge losses of a few major erythrocyte membrane proteins such as

29 spectrins, the abundance of the majority of infected erythrocyte membrane proteins remained unchanged during parasite egress (Figure 2C).

Adducin is lost from the host cytoskeleton 10-15 hours prior to parasite egress

In comparing the proteomes of uninfected erythrocytes and infected erythrocytes, we observed that a subset of host proteins identified with high confidence in the uninfected membrane samples were absent or highly reduced (>>3 fold change) in both the infected erythrocyte and post-rupture vesicle membranes (Table 1), suggesting that major cytoskeletal modifications may be occurring far earlier in the erythrocytic life cycle.

We observed that erythrocyte adducins, tropomyosin and Rac1 disappear from the membrane fraction around 35 hpi, which was striking as these proteins have been shown to be important for the integrity of the cytoskeleton (Kalfa et al., 2006). Direct loss of adducin via mutations or through the loss of Rac1/2 has been shown to disrupt spectrin/actin junctions,(Kalfa et al., 2006) leading to spherocytosis and increased hemolysis in in vivo mouse models (Gilligan et al., 1999; Robledo et al., 2008). Western blot analysis of infected erythrocyte membrane fractions confirmed that a-adducin is present through the first two-thirds of the intraerythocytic life cycle but is lost from the host cytoskeleton at ~35 hpi (Figure 3A). These data was corroborated using immunofluorescence, which showed that adducin is localized to the plasma membrane of uninfected and early stage infected erythrocytes (up to 30 hpi), but disappeared around 35 hpi (Figure 3B). Uninfected erythrocytes also showed an abundance of cytoplasmic adducin staining, which is diminished in infected cells possibly due to parasite digestion of host cytoplasm. We also show that adducin loss from the cytoskeleton is not dependent on calpain, as calpastatin loading into hypotonically lysed and resealed erythrocytes did not prevent adducin loss from the membrane fractions of 30 infected erythrocytes by western blot analysis (Figure 3C). The loss of a-adducin, as well as tropomyosin and Rac1 following schizogony suggests that cytoskeletal dismantling in preparation for egress begins much earlier than previously thought.

Analysis of proteome changes reveals global proteolysis of the host cytoskeleton during parasite egress

Considering the previously proposed role of proteases in parasite egress, we next focused our analysis of the proteomic data on proteins that appeared to be proteolyzed during egress. To do this, we searched for proteins that appeared at significantly smaller sizes than expected in the post-rupture vesicle membrane proteome compared to the infected erythrocyte and uninfected erythrocyte proteomes, as they likely represented proteins proteolytically processed during parasite egress. We used several overlapping methods to determine proteins that appeared smaller in size. Spectral counts for each protein were plotted against their position in the actual gel lane to create

“chromatograms” representing their migration pattern in the SDS-PAGE gel (Figure 4A) for each proteome, which provided a simple yet efficient tool to compare progressive changes in protein distribution patterns between the three proteomes. Although cytoskeletal proteins such as a-spectrin, ankyrin, actin, and band 4.1 were identified at their expected molecular weight in the infected erythrocyte sample, these proteins were also detected at lower molecular weight areas of the chromatogram in the post-rupture vesicle membrane samples, suggesting that they had been proteolyzed. It is worth noting that ankyrin had been previously shown to be proteolyzed by crude parasite lysates (Raphael et al., 2000). However, the majority of proteins, such as flotillin-1 and -

2, showed no apparent cleavage, as shown by the lack of any low-molecular weight fragments (Figure 4A, top panel). This revealed that although several proteins from the 31 post-rupture vesicle membrane proteome appeared smaller than their predicted size

(Figure 4A, lower panel), the majority of proteins did not undergo any proteolytic cleavage during rupture and appeared at their expected molecular weight in all three proteome samples.

Using this method, we estimated the number of proteolyzed proteins in all three proteomes. We observed a doubling in the number of proteolyzed proteins (2% to 5%) from uninfected to infected erythrocyte membrane proteome, however, the magnitude of proteolysis was significantly increased in the post-rupture vesicle proteome (12%)

(Figure 4B). Proteins that had undergone proteolysis were then classified into functional categories: the majority of these proteins (approximately 60%) were associated with or members of the cytoskeleton (Figure 4C). To date, most of these cytoskeletal proteins have not been reported to be proteolyzed in uninfected erythrocytes or during P. falciparum infection.

To confirm our predictions, we conducted western blot analysis of potential proteolyzed hits over a timecourse throughout the parasite life cycle (10-hour increments from 10-40 hpi), in comparison to uninfected erythrocytes (U) and post-rupture vesicles

(R). Processing of several proteins at the time of rupture that were found to be proteolyzed from the MS data were confirmed, including the key cytoskeletal proteins

α/β-spectrins and ankyrin-1 (Figure 5A), as fragments appeared in the post-rupture lane that were similar in size to those found in the MS analysis (see Figure 4A). Several lipid raft-associated proteins including the flotillins were protected from proteolysis, and served as loading controls for the rest of the western blots shown in Figure 5A.

We also identified the presence of several proteases in the infected erythrocyte proteome (Table 2). Notable among these was host calpain-1 that was present only in the late-stage infected erythrocyte membranes and post-rupture vesicle membranes.

Calpain-1 activation and its corresponding association with the erythrocyte membrane are important for malaria parasite egress, as we described recently (Chandramohanadas et al., 2009). We investigated whether calpain activity could account for the degradation of much of the observed cytoskeletal proteolysis during parasite egress, via incubation with erythrocyte membrane fractions in the presence of Ca2+ for 30 minutes. We provide in vitro data validating that, indeed, activated calpain-1 can cleave a whole spectrum of cytoskeletal proteins, generating fragments of similar size to those found in our MS and western blot analyses of post-rupture vesicles (Figure 5B). In addition to calpain-1, parasite-derived SERA proteins, which have been postulated to have a role in egress, were abundantly found in our infected erythrocyte and post-rupture vesicle membrane samples (Yeoh et al., 2007).

Discussion

Our proteomic data revealed that several host proteins, including most prominently

α/β-adducin, disappeared from the host erythrocyte cytoskeleton significantly prior to parasite egress. Targeted disruption of adducin genes has been long known to result increased fragility of the erythrocyte membrane that ultimately leads to spherocytosis and increased hemolysis in mice (Chen et al., 2007; Gilligan et al., 1999; Robledo et al.,

2008). Thus this initial weakening of the cytoskeleton via the loss of the adducins

(perhaps the loss of tropomyosin and Rac1 as well) may be required for the later destruction of membrane integrity to permit facile parasite escape. We determined that

α-adducin loss from the erythrocyte cytoskeleton is both temporally and mechanistically 33 distinct from the changes enacted on the host cell at the time of egress by host calpain-

1. This suggests a new mechanism used by malaria parasites, occurring 15-20 hours prior to parasite egress in order to prepare the infected erythrocyte for rupture. This process may be mediated by parasite kinases to modify host proteins during egress since phosphorylation of cytoskeletal proteins has been shown to result in progressive changes to the cytoskeletal network (Lu et al., 1985; Manno et al., 1995; Soong et al.,

1987). Though parasite kinases have been shown to be exported into the host cell

(Nunes et al., 2007; Vaid et al., 2010), a direct link between parasite kinase activity and infected erythrocyte cytoskeletal changes during egress has yet to be explored.

In addition to the loss of α-adducin, our proteomics data revealed that a significant proportion of erythrocyte cytoskeletal proteins were proteolyzed upon parasite egress. In fact, in post-rupture vesicle membranes, the total amount of spectrins diminished to

~10% of that found in uninfected erythrocyte membranes. In total, more than 60% of the proteins that were found to be proteolyzed were associated with the host cytoskeleton including most prominently the spectrins, ankyrin, and actin, suggesting that global and pathological proteolysis of the host cytoskeleton occurs during the parasite egress process (~48 hpi). Several proteases were also found associated with the infected erythrocyte and post-rupture vesicle membranes, including host calpain-1, an essential enzyme that apicomplexan parasites utilize for egress (Chandramohanadas et al.,

2009). We show that calpain-1 can proteolyze most cytoskeletal proteins during egress, which likely accounts for the striking breakdown of the host erythrocyte membrane around the parasite. Although calpain-1 is capable of proteolyzing many of the cytoskeletal proteins, it is possible that other putative parasite proteases such as SERA family proteases can also perform this function; importantly, SERA5/6 were found to be

34 in abundance on infected erythrocyte membranes and post-rupture vesicles. Although there is little data concerning the proteolytic activity of SERA proteins or for their direct role during parasite egress from the erythrocyte, it is also possible that they contribute to the egress process through destabilization of the host membrane via a non-proteolytic role.

Table 1

Table 1. Proteins that are absent/reduced in infected erythrocyte and/or post-rupture vesicle membranes. List of high abundance proteins that were identified in the uninfected erythrocyte membranes and absent or reduced in the infected erythrocyte and post-rupture vesicle membrane samples.

Table 2A. Proteases identified in the proteomic analyses from infected erythrocyte membranes (I).

Table 2B. Proteases identified in the proteomic analyses from post-rupture vesicle membranes (R).

Table 2. Proteases identified from infected erythrocyte and post-rupture vesicle membrane samples. A list of proteases identified with four or more unique peptides from (A) infected and (B) post-rupture vesicle membrane proteomes are listed.

Figure 1. Fractionation strategy for comparing uninfected erythrocyte, infected erythrocyte and post-rupture vesicle membrane proteomes.

To understand the global changes in the erythrocyte membrane proteome upon parasite egress, parasite- infected (I) and post-rupture vesicle membranes (R) were isolated and compared with uninfected erythrocyte membranes (U). Newly replicated parasites (P) and the food vacuole following schizogony (FV) are also shown. (A). Uninfected erythrocyte membranes were prepared by hypotonic lysis and ultracentrifugation. The infected erythrocyte membrane proteome was prepared by saponin treatment of magnet-purified 37

schizonts and ultracentrifugation. To prepare post-rupture vesicle membranes, we used a differential centrifugation strategy to purify vesicles from media following parasite egress (see methods for more details). All the samples were solubilized in Laemmli buffer, resolved by SDS-PAGE and visualized by colloidal Coomassie staining. 34 slices were excised from each proteome for analysis by mass spectrometry. (B). To illustrate the efficiency of the fractionation strategy, the purification of post-rupture vesicles was visualized by microscopy. Each fraction was stained with FM4-64 for membrane staining and DAPI for DNA. Panel 1: Uninfected erythrocytes; Panel 2: crude media just after parasite rupture from infected erythrocytes containing merozoites (arrow marks), post-rupture vesicles (arrowheads) and intact erythrocytes (broken line); Panel 3: purified post-rupture vesicles derived from parasite egress. (C). Selective enrichment of infected erythrocyte membranes from parasite material was also confirmed by western blot analysis using a transgenic parasite line expressing cytosolic GFP. Protein from whole infected erythrocytes (I), purified parasite cells fractionated by low speed centrifugation from saponin-treated infected erythrocytes (P), or post-rupture vesicles fractionated from media after parasite egress (R) were separated by SDS-PAGE. Efficient isolation of infected erythrocyte membrane material was determined by western using a parasite cytoplasmic marker (anti-GFP) and a human erythrocyte membrane marker (anti-Band 4.9).

Figure 2. Comparative proteomic analysis of the erythrocyte membrane proteome during parasite egress.

(A). Infected erythrocyte membranes (I) and post-rupture vesicles derived from parasite egress (R) were isolated and compared with uninfected erythrocyte membranes (U). Samples for the proteomic analysis were resolved on a 4-12% SDS-PAGE gel and stained with colloidal Coomassie. 34 slices were excised from each sample lane, digested with trypsin, extracted, and analyzed by LC/MS/MS mass spectrometry- based sequencing. The overall migration pattern for the post-rupture vesicle membrane proteome was 39 different, as the characteristic high molecular weight spectrin doublet appeared to be degraded and with the appearance of more small molecular weight bands. (B). Peptides from each gel slice were sequenced by mass spectrometry and the percentage distribution of identified proteins through the SDS-PAGE lanes for all samples (U, I, and R) were expressed in a heat map format. Red color indicates high percentage of peptides per protein (10%) while yellow indicates minimum (0%). This representation indicates that proteins were well distributed throughout the SDS-PAGE gel. (C). Spectral counting was used to compare the relative abundance of each protein from the three proteomes. The total number of peptides was summed for each identified protein in the infected erythrocyte and post-rupture vesicle proteome samples and was individually compared to the uninfected preparation to determine fold changes in relative protein abundance among the three proteomes. The abundance of most proteins remained similar (within a 3-fold difference) among the three proteomes, whereas a small proportion (~10%) showed markedly lower abundance in the post-rupture vesicle proteome.

Figure 3. Loss of adducin from the erythrocyte cytoskeleton prior to

parasite escape is calpain-independent.

oaded oaded l - n invasion. -

independent. -

Western blot analysis of calpastati

(C).

infected erythrocytes. -

Immunofluorescence confirms the presence of adducin in uninfected, ring, and

(B).

confirms that the loss of adducin from the infected erythrocyte cytoskeleton around 35 hpi is calpain is hpi 35 around cytoskeleton erythrocyte infected the from adducin of loss the that confirms

levels are shown as a loading control.

Western blot analysis confirmed the loss of adducin from the membrane fraction of infected erythrocytes by 35 hours post illin (A). Flot trophozoite infected erythrocytes, and infected its erythrocytes absence in schizont

Figure 4. Proteomic profiling shows extensive cytoskeletal protein proteolysis during parasite egress.

(A). To determine proteins that were proteolytically processed, the spectral count for each protein was plotted against the corresponding gel slice number (and hence the apparent molecular weight) judged from its migration in the SDS-PAGE gel. Several proteins appeared smaller than their expected molecular weight as indicated by a second peak in the chromatogram corresponding to a lower molecular weight as seen in α- spectrin, band 4.1 and actin. Abundance of the smaller species was universally increased in the post-rupture 42 vesicle proteome sample. Otherwise, the majority of the proteins showed no major difference in abundance and molecular weight as exemplified by flotillin-1. (B). Proteolysis of host membrane proteins significantly increases at egress. Number of processed proteins identified from the uninfected (U), infected erythrocyte (I) and post-rupture vesicle (R) membrane proteomes is expressed as a percentage of total proteins identified. (C). Proteins that were processed in the post-rupture vesicle erythrocyte membrane were classified according to their functions as obtained from the human proteins reference database (www.hprd.org), and corresponding Gene Ontology (GO) annotation.

Figure 5. Validation of calpain-dependent cytoskeletal processing during parasite rupture.

(A). Western blot analysis from uninfected erythrocyte (U), infected erythrocyte throughout the life cycle (10- 40 hpi), and post-rupture vesicle (R) membranes was used to validate the proteolytic processing of candidate proteins. Analysis of α-spectrin, β-spectrin and ankyrin-1 showed major degradation of target proteins only in the post-rupture vesicle proteome sample whereas many proteins, such as flotillin (shown as loading controls underneath each blot) showed no proteolytic processing. (B). Erythrocyte membranes treated for 30 min with activated human calpain-1 were able to recapitulate the processing observed in post- rupture vesicles. Western blot analysis of calpain-mediated processing is shown for α-spectrin, β-spectrin, and ankyrin-1.

Figure 6. New model for parasite driven modifications to the host cytoskeleton during parasite egress.

the formation of large holes

mediated proteolysis of the remaining cytoskeleton at 48 hpi to eventually - scale scale calpain -

We proposethat parasite egress begins as earlyas 35 hpiwith the lossof spectrin/actinadaptors, and lead to in the cytoskeleton mesh, followed release by merozoites. large

IV. Host GPCR signaling necessary for parasite-mediated cytolysis: Thwarting host signaling to stop them in their tracks Parasite-induced host cell cytolysis has been suggested to be a two-step, Ca2+- dependent process (Salmon et al., 2001; Wickham et al., 2003b) requiring both host cell ion loss (Moudy et al., 2001) and increased membrane poration (Abkarian et al., 2011;

Glushakova et al., 2010). Recent reports implicate the activity of parasite-derived proteins including: a perforin-like protein (Kafsack et al., 2009), a kinase (Dvorin et al.,

2010), and proteases (Arastu-Kapur et al., 2008; Yeoh et al., 2007) though reports of host protein contribution to this process are limited. We have recently shown that the host-derived protease calpain is required for exit of both P. falciparum and T. gondii and functions to proteolyze the actin cytoskeleton just prior to cytolysis (Chandramohanadas et al., 2009; Millholland et al., 2011). Furthermore, we have identified calpain- independent loss of adducin from the host erythrocyte actin cytoskeleton prior to P. falciparum exit (Millholland et al., 2011). Given the temporal separation of these striking changes to the host cytoskeleton during the last phase of the intracellular cycle, we hypothesized that a complex host pathway may connect these changes and result in the disintegration of the host plasma membrane. Identification of host proteins necessary for parasite infection may offer an untapped resource of antiparasitic targets.

Host RNAi screen reveals a pathway necessary for T. gondii-mediated cytolysis

To identify host genes essential for parasite-mediated cytolysis, we performed an

RNAi screen in U2OS cells (highly transfectable T. gondii host cells) focused on canonical Ca2+-signaling components, given our earlier studies that implicated host calpain in T. gondii and P. falciparum exit (Chandramohanadas et al., 2009, Millholland et al., 2011). To circumvent the potential for functional redundancy between host gene

46 isoforms, we tested pooled siRNAs for simultaneous knockdown of gene families, as well as individual genes present in both U2OS cells and erythrocytes. This primary screen included 45 individual gene knockdowns and 11 multi-isoform knockdowns

(Tables 3,4). Pooled siRNAs (three siRNAs/gene) were arrayed in quadruplicate, reverse-transfected into U2OS cells, and infected 24 hours post-transfection with transgenic T. gondii tachyzoites (MOI of 0.1) that constitutively secrete GFP into the PV space, allowing for facile identification of parasite doublings as a function of life cycle progression (P30-GFP; Figure 7A, Figure 8; (Striepen et al., 1998). siRNAs against calpain small subunit (capns1) and a scrambled oligo (Scr) were included as positive and negative controls, respectively. Knockdown efficiency was maximal at 72 hours post-transfection, which corresponded with parasite-mediated cytolysis in this cell type

(~50 hpi). Following synchronous T. gondii infection (Kafsack et al., 2004), plates were fixed and imaged at 6, 24, 44, or 60 hpi to assess parasite life cycle progression through cytolysis. Genes involved in parasite-mediated cytolysis showed an accumulation of PVs containing >64 parasites at 60 hpi upon knockdown (Z score +1.5, p < 0.05), ~10 hours after host cell rupture typically occurs in this cell type.

From this screen, we initially identified 5 gene families whose knockdown blocked parasite-mediated host cell cytolysis: calmodulin (CaM), Ca2+/calmodulin dependent kinases (CaMK), Gα subunits (Gα), phospholipase C (PLC) and protein kinase C (PKC)

(Figure 7B). To deconvolute the specific genes required for parasite-mediated cytolysis from these gene family hits, we tested two unique siRNAs targeting individual and pairs of genes (Figure 7C, Table 5). We determined that single knockdown of gnaq

(p<0.002), and knockdown of the gene pairs prkca/prkcb (p<0.002), plcb1/plcg1

(p<0.002), calm1/calm2 (p<0.05) and camk1/camk2a (p<0.05), showed a statistically

47 significant and correlated accumulation of unruptured PVs at 60 hpi (r=0.91; Figure 7D).

Stable shRNA-mediated knockdown of these genes had no effect on parasite doublings, reinforcing their function in cytolysis rather than parasite replication (Figure 7E). shRNA knockdown efficiency using multiple oligos was measured by western blot analysis of total protein as compared to scr (Figure 7F). Representative images are shown for shRNA knockdown cell lines that resulted in an accumulation of unruptured PVs in comparison to controls (scr) or negatives (prkaca/prkacb) that show newly reinvaded parasites at 60 hpi (Figure 7G).

Immunodepletion studies indicate conserved host protein function in P. falciparum-mediated cytolysis

We next investigated whether the protein products of the validated gene hits in T. gondii functioned similarly in P. falciparum-induced erythrocyte cytolysis, via antibody- mediated depletion studies. To do this, erythrocytes were hypotonically lysed prior to incubation with target antibodies preconjugated to sepharose beads. Removal of the target protein-antibody conjugates were achieved via centrifugation and manipulated erythrocytes were hypertonically resealed prior to P. falciparum infection. Efficiency of target protein immunodepletion in erythrocytes was determined by western blot and only immunodepletions showing >80% knockdown were further analyzed (Figure 9A;

(Chandramohanadas et al., 2009). Synchronous P. falciparum parasites were assessed for life cycle progression by Giemsa smear (Figure 9B) and flow cytometry of DNA content (Figure 9C; Figure 10) to distinguish blocked multinucleated schizonts from newly invaded, ring-form parasites with a single nucleus. Immunodepletion of Gαq,

PLCβ1, CaM-1, or the simultaneous immunodepletion of both PKCα and PKCβ resulted in an accumulation of schizont stage parasites and a lack of newly invaded rings at 60

48 hpi, indicating a block in parasite-mediated cytolysis and corroborating the results of our

RNAi screen in T. gondii host cells. To assess parasite viability, trapped parasites were mechanically released from depleted host cells and assessed for their invasive capacity via flow cytometric quantitation of mononuclear ring-stage parasites 12 hours following mechanical release (Figure 9D). Parasites trapped within erythrocytes were able to recover invasive capacity upon needle-shearing only up to 54 hpi, indicating that parasite death likely occurs following 6 hours of blocked cytolysis.

Host PKC is Activated During Schizogony to Remove Adducin from the Host

Cytoskeleton

As PKC is a major downstream effector of Gαq-coupled GPCR signaling via PLC- mediated generation of diacylglycerol (DAG; (Castagna et al., 1982; Rhee et al., 1989) and PKC inhibitors showed potent antiparasitic activity and the ability to block parasite- mediated cytolysis in vitro (Figure 12A), we were interested in understanding the role of this critical host enzyme in parasite life cycle progression. To measure host PKC activity during the P. falciparum life cycle, we generated a transgenic P. falciparum line that secretes a FRET-based PKC activity indicator (CKAR; (Violin et al., 2003) into the infected erythrocyte cytoplasm (3D7-CKAR) via expression of a chimeric construct with an N-terminal signal peptide and PEXEL motif (Gallegos et al., 2006; Hiller et al., 2004;

Marti et al., 2004; Violin et al., 2003). Figure 11A displays the specificity of the CKAR reporter for PKC activity with representative positive control FRET images (maximal

FRET signal) of 3D7-CKAR-infected erythrocytes treated with the PKC agonist phorbol myristate acetate (PMA) and negative control images (minimal FRET signal) upon treatment with the PKC inhibitor Gö6976 or PKC immunodepletion prior to infection. As with any protein present in the erythrocyte cytoplasm, a significant amount of CKAR was

49 taken up into the parasite digestive vacuole, causing an intense CKAR FRET signal within this organelle likely due to intermolecular FRET. In order to account for this artifact in our measurements of host PKC activity, 3D7-CKAR parasites were removed from their host cells and intracellular FRET signal was quantified to assess this background signal within the digestive vacuole (Figure 12C). By FRET microscopy (Figure 11B) and background-corrected fluorometry to remove digestive vacuole signal (Figure 11C), we show limited host PKC activity throughout the ring and trophozoite stages (0-25 hpi) with a sharp increase during schizogony (30-40 hpi). Immunodepletion of Gαq or PLCβ1 ablates PKC activity during the entire infectious cycle, reinforcing the importance of the host Gαq signaling pathway upstream of PKC activation (Figure 12D). PMA restored

PKC activity in these parasite-infected erythrocytes depleted of signaling components and also rescued the associated exit defect (Figure 12E), further indicating a critical role for host PKC. PKC activity reporter cell lines were also generated via stable expression of CKAR in U2OS host cells prior to T. gondii infection. The CKAR signal output throughout the T. gondii life cycle in these reporter U2OS cells largely mirrored the results obtained in P. falciparum, with maximal FRET signal occurring in the last third of the intracellular life cycle (Figure 11D-F). Knockdown of host gnaq or plcb1/plcg1 ablates PKC activity but this activity was restored by the PKC agonist PMA (Figure

11H,I).

In a previous chapter, we have shown that the host cytoskeletal protein adducin is lost from the erythrocyte actin cytoskeleton at ~35 hpi of P. falciparum in a calpain- independent manner (Millholland et al., 2011). As adducin cytoskeletal association is regulated by the PKC-mediated phosphorylation of residues S716/S726 (Matsuoka et al., 1996), we assessed PKC depletion on host adducin cytoskeletal association during

50 parasite infection. Western blot and immunofluorescence analysis of host cytoskeletal fractions confirms adducin disappearance from both P. falciparum-infected erythrocytes

(Figure 11G, top; Figure 12F) and T. gondii-infected U2OS cells late in the intracellular cycle (Figure 11H, top; Figure 12G). However, PKCα/β immunodepletion from erythrocytes (Figure 11G, bottom) or shRNA knockdown in U2OS cells (Figure 11H, bottom) abrogates this loss and maintains adducin cytoskeletal association through the end of both parasite life cycles. The S716/S726 phosphorylated adducin species (p- adducin) accumulates in mock-treated cells in the last third of the life cycle by western blot while the unphosphorylated form is lost from cytoskeletal fractions, unless PKCα/β is depleted from host cells. Gαq or PLCβ1 immunodepletion from erythrocytes (Figure

12F), or knockdown in U2OS cells (Figure 12G) also caused adducin persistence within the host cytoskeleton, highlighting the function of this upstream cascade.

In order to examine the effect of adducin persistence on T. gondii life cycle progression, we overexpressed the PKC phospho-mutant adducin (S716A/S726A;

(Matsuoka et al., 1998) as well as wild-type adducin in host A549 cells. S716A/S726A adducin expression resulted in minimal phosphorylation by western blot upon T. gondii infection (Figure 11I), while wild-type adducin overexpressors show an abundance of p- adducin. Similar to PKCα/β stable knockdown, S716A/S726A adducin expression resulted in the persistence of enlarged vacuoles by 60 hpi (Figure 11J,K), with no effect on parasite replication (Figure 11L), suggesting that PKC-induced adducin loss directly mediates host cell cytolysis. Tachyzoites trapped within host cells show reduced viability following 18-24 hours of blocked cytolysis within host cells expressing the adducin phospho-mutant S716A/S726A, similar to PKC inhibition or knockdown (Figure 12B),

51 highlighting the feasibility of targeting this phase of the parasite life cycle as an antiparasitic strategy.

Host signaling cascade is required for calpain-mediated cytoskeletal proteolysis

Host calpain activation has been shown to result in cytoskeletal proteolysis necessary for parasite exit (Chandramohanadas et al., 2009; Millholland et al., 2011).

We thus assessed calpain activation following immunodepletion of signaling components from erythrocytes using an activity-based probe for cysteine proteases, DCG04 (Figure

13A; (Greenbaum et al., 2000). As opposed to negative control PKAα/β immunodepleted cells which show a clear active calpain band at the conclusion of the life cycle, PKCα/β, PLCβ1, and Gαq immunodepletion disallows calpain activation, similar to calpain immunodepleted cells. CaM-1 immunodepletion resulted in diminished calpain activation, indicating that CaM-1 may facilitate this process. As a corollary to calpain activation, we examined spectrin cleavage via western blot, a key calpain substrate. Immunodepletion of pathway components abrogates spectrin proteolysis in P. falciparum-infected erythrocytes at 50 hpi, while PKAα/β-immunodepleted cells show multiple spectrin fragments (Figure 13B). Interestingly, in vitro incubation of U2OS cell membrane fractions with activated CaMKII prior to incubation with calpain resulted in multiple spectrin cleavage products (Figure 13C), suggesting that CaMKII phosphorylation of cytoskeletal substrates may enhance calpain-mediated proteolysis.

CaMKII activation is shown by CaM-1 co-immunoprecipitation and autophosphorylation by p-CaMKII western blot analysis near the end of the T. gondii life cycle (Figure 13D), suggesting that host CaMKII activity may enhance cytoskeletal proteolysis to facilitate parasite release.

Analysis of host Ca2+ dynamics Implicates TRPC6 in Ca2+ influx 52

Finally, we determined changes in host [Ca2+] that drive this network. To assess erythrocyte [Ca2+] during P. falciparum infection, we confined a Ca2+ indicator to the erythrocyte space via loading of a fura-2-dextran conjugate. Within the U2OS cell cytoplasm during T. gondii infection, Ca2+ measurements were achieved via expression of the ratiometric FRET-based Ca2+ indicator yellow cameleon 3.6 (YC3.6) in U2OS cells (Nagai et al., 2004; Palmer and Tsien, 2006). By microscopy and background- corrected fluorometry (Figure 14A,B), we show the relative changes in host cell [Ca2+] to be similar between both parasite systems, with host [Ca2+] hovering near resting levels

(100 nM) throughout the first two-thirds of the life cycle, followed by a sharp increase in

[Ca2+] at the point of parasite release.

Given the large [Ca2+] increase seen in both T. gondii and P. falciparum host cells, we further hypothesized that host plasma membrane cation channels play a role in this influx. siRNA-mediated knockdown studies of mechanosensitive transient receptor potential (TRP) channels (trpc1, trpc3, trpc6) as well as canonical endoplasmic reticulum

Ca2+ channels (ryr3, ip3r1) identified TRPC6 as a specific mediator of T. gondii exit, as trpc6 knockdown resulted in an accumulation of unruptured vacuoles by 60 hpi (p<0.05;

Figure 14C,D). Trpc6 knockdown using multiple siRNA oligos abrogated the large increase in cytoplasmic [Ca2+] while ip3r1 knockdown diminished the initial minor increase in [Ca2+] observed earlier in T. gondii infection, indicating that PLC activity and generation of IP3 may be responsible for the minor increase in [Ca2+] seen only in T. gondii-infected cells (Figure 14E).

Pharmacological studies in P. falciparum-infected erythrocytes confirmed the importance of mechanosensitive cation channel function in parasite-mediated cytolysis

2+ (Figure 15A). [Ca ] diminished in cells depleted of Gαq, PLCβ, or PKCα/β (Figure

15B,C), but was rescued by PKC activation with PMA in cells depleted of Gαq or PLCβ, indicating that PKC activity facilitates later Ca2+ influx, perhaps via adducing phosphorylation and loss. PMA could not rescue Ca2+ influx in cells depleted of PKCα/β, further confirming the necessity of PKC activity for this process. Taken together, these data highlight the importance of TRPC6-mediated cation influx in host cell cytolysis, perhaps as a mechanosensitive response to cytoskeletal rearrangement.

Mammalian PKC inhibitor demonstrates anti-parasitic activity in vivo

In order to assess the importance of this signaling cascade in parasitic disease progression, we endeavored to test inhibition of this cascade on parasite burden in vivo.

Given that PKC has been targeted for drug development efforts in multiple disease contexts from cancer to transplant rejection, and there are no known orthologs of the classical PKC enzymes in apicomplexan genomes, we undertook studies of known PKC inhibitors in murine models of malaria and toxoplasmosis. Intravenous injection of 10 mg/kg Gö6976, an inhibitor of classical mammalian PKC enzymes, caused a significant decrease in parasitemia in a mouse P. yoelii malaria model in the classic 4-day Peter’s suppression test (Figure 16A). Similarly, intraperitoneal injection of 10 mg/kg Gö6976 limited T. gondii parasite burden in both the spleen (p<0.0001; Figure 16B, top) and peritoneal exudate cells (p<0.0001; Figure 16B, bottom). The consistency in antiparasitic activity in vivo further suggests the importance of host PKC function in both parasitic infections and underscores the conserved function of this host-signaling pathway.

As further corroboration of the central role of host PKC in severe malaria, we undertook studies of the orally-bioavailable specific PKC inhibitor sotrastaurin in a mouse model of experimental cerebral malaria (Figure 16C,D). This inhibitor has 54 passed Phase I trials and is undergoing Phase II trials for numerous indications (Budde et al., 2010; Friman et al., 2011; Wagner et al., 2009), underscoring its potential as an antimalarial drug candidate. 50 mg/kg by gastric gavage in a standard 4-day Peters’ suppression test led to a significant decrease in P. berghei ANKA parasitemia

(p<0.0001; Figure 16C) and significantly increased survival versus vehicle-treated controls (p=0.0029; Figure 16D). We show direct translation of biological discovery to therapeutic design: identification of this host signaling activity translates directly to a drug discovery effort, as known inhibitors of extensively-studied host PKC enzymes control parasitic disease in vivo. Given the multitude of other active proteins within the cascade, it is clear that these newly identified host mediators of parasite-induced cytolysis represent an untapped resource of antiparasitic targets.

TCA metabolites may initiate the cytolytic cascade through interaction with

Gα q-coupled GPCRs

As Gαq, PLC, and PKC converge in established GPCR signaling networks, we hypothesized that GPCRs may play a role in initiation of this host pathway. Considering that parasites do not encode GPCRs, we suggest a parasite-derived small molecule ligand may signal to GPCRs of host origin through direct interaction with the host cell membrane to mediate signaling through Gαq. We found it striking that several parasite metabolites peak during schizogony, especially the TCA cycle intermediate alphaketoglutarate (αKG; (Olszewski et al., 2009). As αKG and other TCA cycle intermediates including succinate have recently been shown to signal through Gαq- coupled GPCRs, such as oxoglutarate receptor 1 (oxgr1) and succinate receptor 1

(sucnr1; (He et al., 2004; Qi et al., 2004), we hypothesized that increased parasite metabolism during periods of high replication may initiate host cell signaling leading to

55 cytolysis. To test this idea, we first confirmed by western blot analysis that OXGR1 and

SUCNR1 were found in erythrocytes (Figure 17A) and U2OS cells (Figure 17B) and not in parasites. Immunofluorescence studies of both GPCRs revealed puncta on the host plasma membrane in uninfected and infected cells; however, there was a surprising enrichment on the PV membrane in both P. falciparum-infected erythrocytes and T. gondii-infected U2OS cells. To test whether these host GPCRs play a role in the signaling cascade, we undertook knockdown studies of metabolite-sensing GPCRs in

U2OS cells and assayed for delays in T. gondii-mediated cytolysis. Simultaneous knockdown of oxgr1 and sucnr1 caused an accumulation of large PVs at 60 hpi (p<0.09) versus negative control cells or knockdown of the Gαi–coupled lactate receptor gpr81, in which cytolysis occurred by 52 hpi (Figure 17C, D).

To further validate the knockdown results, we assessed whether activation of these host GPCRs occurs prior to parasite exit. To do this, we utilized commercially available Chem-1 cell lines (Millipore) that allow direct measurement of specific GPCR activity through overexpression of Gαq and only OXGR1 or SUCNR1, on a parent cell line background which expresses few endogenous GPCRs. T. gondii infection of Chem-

1 cells overexpressing either OXGR1 or SUCNR1 showed GPCR activity (as measured by Fluo-8 fluorescence) to persist during the last third of the parasite life cycle until cytolysis (Figure 17E, F). Conversely, the parental Chem-1 cell line displayed limited

GPCR activity throughout the T. gondii life cycle and a significant delay in parasite- mediated cytolysis (Figure 17G), further suggesting a role for metabolite-sensing

GPCRs. Satisfyingly, a mixture of exogenous αKG/succinate initiated a hastening of T. gondii exit, occurring ~20 hours sooner than incubation with DMSO alone or lactate

(Figure 17H). Furthermore, αKG/succinate induced premature lysis of P. falciparum parasites leading to parasite death (Figure 17I).

Discussion

Host cell cytolysis was thought to be largely parasite-mediated, mainly due to increasing pressure on the host cell membrane and cytoskeleton by the growing parasite body (Glushakova et al., 2010; Glushakova et al., 2005). However, we have recently shown that host calpain-mediated proteolysis of host cytoskeletal proteins is necessary for parasite exit (Chandramohanadas et al., 2009; Millholland et al., 2011). In this work we present a complex host-derived signaling pathway that functions in parasite- mediated cytolysis, suggesting that host cell cytolysis is a highly regulated process requiring a complex interplay of host-derived components.

We present a model (Figure 18) in which diffusion of parasite-derived small molecule GPCR ligands outside into the PV space and host cell results in overstimulation of host GPCRs. These GPCRs engage a Gαq-mediated signaling pathway converging on PKC activation that results in the complete loss of a key cytoskeletal component, and leads to pathological Ca2+ influx through mechanosensitive plasma membrane TRP channels. Ca2+-charged host CaM activates CaMK which in turn phosphorylates cytoskeletal substrates, increasing their propensity for calpain-mediated proteolysis. Increasing cytoplasmic [Ca2+] induces global calpain activation which results in the proteolysis of various substrates, and in turn causes the loss of host cell plasma membrane integrity to allow for parasite release.

PKC and calpain play key roles within this cascade, a partnership that has been underscored in other human pathologies related to GPCR overstimulation, including glutamate neuroexcitotoxicity and post-myocardial infarction cell death. PKC 57 downregulation has been shown to protect against glutamate neurotoxicity (Ahlemeyer et al., 2002; Favaron et al., 1990; Hasham et al., 1997), while calpain inhibition has been shown to abrogate neuronal cell death (Witt et al., 1994) and block Ca2+ influx (Weiss et al., 1990). Similarly, PKC and calpain are thought to play a role in post-myocardial infarction ischemic cell death of multiple tissues (Bright et al., 2004; Padanilam, 2001;

Piccoletti et al., 1992; Speechly-Dick et al., 1994) as inhibitors of these enzymes offer protective effects (Bevers et al., 2010; Bevers et al., 2009; Koponen et al., 2003; Neuhof et al., 2008; Wagey et al., 2001; Wei et al., 2010). Though it is unclear whether the sequential activation of these enzymes is necessary for cell death in these disease contexts, we suggest that PKC activity is an essential upstream component of pathological calpain activation in the context of parasite infection. Given the antiparasitic activity of PKC inhibitors in mouse models of both toxoplasmosis and malaria, inhibition of multiple host proteins upstream of PKC activity will likely limit parasite growth with low propensity for generation of resistance in the field.

Table 3. siRNA screen in U2OS cells Identifies Host Mediators of T. gondii-Induced Host Cell Cytolysis SINGLE GENES Means shown are representative of at least 3 separate experiments. N=3. Unruptured Unruptured Gene Vacuoles/ SEM Vacuoles/ SEM Full Gene Name GO Term Symbol Field (44 hpi) Field (60 hpi) (44 hpi) (60 hpi) LEGEND: NEGATIVE CONTROL POSITIVE CONTROL

POSITIVE HIT calcium ion binding, calcium- anxa1 annexin A1 1.53 0.244 0.0427 0.0295 dependent phospholipid binding, phospholipase inhibitor activity

59 calcium ion binding, calcium- dependent phospholipid binding, anxa2 annexin A2 1.07 0.332 0.0632 0.0552 phospholipase inhibitor activity cytoskeletal protein binding ATPase, Ca++ transporting, calcium ion transport, cation atp2a3 1.74 0.188 0.0222 0.0329 ubiquitous transport ion transmembrane transport, ATPase, aminophospholipid atp8b1 1.432 0.182 0.0341 0.0328 phospholipid transport, membrane transporter, class I, type 8B, member fraction cell surface receptor linked signaling cabin1 calcineurin binding protein 1 1.69 0.152 0.0281 0.0388 pathway plasma membrane, G-protein calcr calcitonin receptor 1.14 0.127 0.0733 0.0621 signaling, coupled to cAMP nucleotide second messenger calcium ion binding, calcium- calm1 Calmodulin 1 1.94 0.159 0.147 0.0913 dependent protein binding calcium ion binding, calcium- calm2 Calmodulin 2 1.32 0.273 0.0986 0.0325 dependent protein binding calcium/calmodulin-dependent camk1 1.09 0.299 0.114 0.0374 signal transduction protein kinase I calcium- and calmodulin-dependent calcium/calmodulin-dependent camk2a 1.53 0.263 0.195 0.0522 protein kinase complex; calmodulin protein kinase II alpha binding calcium/calmodulin-dependent calcium ion binding, calmodulin camk2b 0.978 0.264 0.166 0.0426 protein kinase II beta binding calcium ion binding, calcium- capn1 calpain 1, (mu/I) large subunit 1.42 0.283 0.522 0.0893 dependent cysteine-type endopeptidase activity calcium ion binding, calcium- capn2 calpain 2, (m/II) large subunit 1.19 0.174 0.476 0.0783 dependent cysteine-type endopeptidase activity signal transduction, calcium- capn5 calpain 5 1.57 0.293 0.156 0.108 dependent cysteine-type endopeptidase activity calcium ion binding, calcium- capns1 calpain, small subunit 1 1.21 0.273 1.21 0.214 dependent cysteine-type 60 endopeptidase activity G-protein coupled receptor protein guanine nucleotide binding protein (G gnaq 1.48 0.284 1.08 0.412 signaling pathway; signal transducer protein), q polypeptide activity Guanine nucleotide binding protein G-protein coupled receptor protein gnai1 (G protein), alpha inhibiting activity 1.324 0.236 0.135 0.0316 signaling pathway; signal transducer polypeptide 1 activity Guanine nucleotide binding protein G-protein coupled receptor protein gnai2 (G protein), alpha inhibiting activity 1.24 0.331 0.0598 0.0566 signaling pathway; signal transducer polypeptide 2 activity Guanine nucleotide binding protein G-protein coupled receptor protein gnai3 (G protein), alpha inhibiting activity 1.05 0.364 0.214 0.0523 signaling pathway; signal transducer polypeptide 3 activity G-protein coupled receptor protein guanine nucleotide binding protein (G gnas 1.34 0.236 0.172 0.0615 signaling pathway; signal transducer protein), alpha s activity G-protein coupled receptor protein guanine nucleotide binding protein (G gna12 1.77 0.164 0.118 0.0427 signaling pathway; signal transducer protein), alpha 12 activity G-protein coupled receptor protein guanine nucleotide binding protein (G gna13 0.96 0.234 0.0464 0.0634 signaling pathway; signal transducer protein), alpha 13 activity potassium intermediate/small plasma membrane, ion transport, kcnn4 conductance calcium-activated 1.24 0.236 0.1953 0.0597 calcium-activated potassium channel channel, subfamily N, member 4 activity intermediate filament, structural lmna lamin A/C 1.05 0.235 0.1655 0.0693 molecule activity mylk myosin light chain kinase 1.09 0.296 0.112 0.0882 kinase activity phosphoinositide-3-kinase, catalytic, pik3ca 1.69 0.253 0.0821 0.0333 signal transduction alpha polypeptide phosphoinositide-3-kinase, catalytic, pik3cb 1.32 0.128 0.184 0.0413 signal transduction beta polypeptide calcium ion binding; calcium- pla2g4a phospholipase A2, group IVA 1.22 0.237 0.142 0.0894 dependent phospholipase A2 activity 61 calcium ion binding, signal plcb1 phospholipase C, beta 1 1.65 0.385 0.121 0.0521 transduction calcium ion binding, signal plcb2 phospholipase C, beta 2 1.21 0.236 0.0541 0.0433 transduction calcium ion binding, signal plcb3 phospholipase C, beta 3 1.55 0.237 0.112 0.0323 transduction calcium ion binding, signal plcd1 phospholipase C, delta 1 1.26 0.235 0.184 0.0166 transduction signal transduction, activation of plcg2 phospholipase C, gamma 2 1.34 0.384 0.193 0.0882 store-operated calcium channel activity calcium ion binding, signal plcg1 phospholipase C, gamma 1 1.31 0.254 0.113 0.063 transduction calcium ion binding, plasma plscr1 phospholipid scramblase 1 1.47 0.295 0.144 0.0511 membrane plasma membrane, cAMP- protein kinase, cAMP-dependent, prkaca 1.01 0.133 0.125 0.0417 dependent protein kinase inhibitor catalytic, alpha activity protein kinase, cAMP-dependent, signal transduction, plasma prkacb 1.08 0.188 0.214 0.135 catalytic, beta membrane signal transduction, plasma prkca protein kinase C, alpha 0.953 0.213 0.242 0.0335 membrane prkcb protein kinase C, beta 1.09 0.223 0.0842 0.0446 signal transduction, plasma membrane prkcg protein kinase C, gamma 1.04 0.192 0.0655 0.0875 signal transduction protein phosphatase 3, catalytic ppp3ca 1.44 0.214 0.0541 0.114 calcium ion transport subunit, alpha isozyme ptk2 PTK2 protein tyrosine kinase 2 0.874 0.122 0.177 0.0552 cytoskeleton, plasma membrane ras-related C3 botulinum toxin plasma membrane, signal rac1 substrate 1 (rho family, small GTP 1.41 0.216 0.0842 0.0224 transduction binding protein Rac1) ras-related C3 botulinum toxin plasma membrane, signal rac2 substrate 2 (rho family, small GTP 1.21 0.177 0.0831 0.0452 transduction binding protein Rac2) plasma membrane, cytoskeleton, rhoa ras homolog gene family, member A 0.915 0.131 0.113 0.0316 signal transduction plasma membrane, signal rhob ras homolog gene family, member B 0.889 0.122 0.145 0.0332 transduction 62 Scr scrambled oligo (negative control) 1.21 0.112 0.0731 0.0221 ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! Table 4. MULTI-ISOFORM FAMILIES AND CONSTITUENTS Means shown are representative of 3 separate experiments. N=3. Unruptured Vacuoles/ Field Unruptured Vacuoles/ Gene Family Single Genes Included SEM (44 hpi) SEM (60 hpi) (44 hpi) Field (60 hpi) LEGEND: NEGATIVE CONTROL POSITIVE CONTROL

POSITIVE HIT anxa1 Annexin 1.19 0.311 0.0594 0.0442 anxa2

Calmodulin calm1

63 0.974 0.223 0.713 0.195 (CaM) calm2

camk1 CaM Kinase camk2a 1.07 0.142 0.822 0.138 camk2b

capn1 Calpain capn2 1.17 0.281 0.973 0.155 capns1

gnaq gnai1 gnai2 G-alpha gnai3 1.11 0.262 0.842 0.155 subunits gnas gna12 gna13

lmna Lamin lmnb1 1.24 0.171 0.121 0.0997 lmnb2

pik3ca PI3-Kinase 0.892 0.174 0.112 0.164 pik3cb

plcb1 plcb2 Phospholipase plcb3 1.22 0.242 0.942 0.232 C plcd1 plcg2 plcg1

prkca Protein Kinase

64 prkcb 1.41 0.331 0.914 0.176 C prkcg

rac1 Rac GTPases 1.09 0.132 0.141 0.0571 rac2

rhoa Rho GTPases 1.11 0.233 0.0912 0.0778 rhob ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! Table 5. GENE PAIRS Means shown are representative of 5 separate experiments. N=5. Gene (s) Unruptured Vacuoles/Field SEM Unruptured Vacuoles/Field SEM (44 hpi) (44 hpi) (60 hpi) (60 hpi) LEGEND: NEGATIVE CONTROL POSITIVE CONTROL POSITIVE HIT Scr #1 1.53 0.244 0.233 0.071 Scr #2 1.07 0.332 0.103 0.099 capn1 #1 1.21 0.212 0.312 0.041 capn1 #2 1.33 0.155 0.441 0.198 capn2 #1 1.13 0.213 0.291 0.171

65 capn2 #2 1.17 0.199 0.212 0.104 capn1 #1/capn2 #1 1.22 0.273 1.31 0.285 capn1 #2/capn2 #2 1.41 0.112 1.05 0.256 capns1 #1 1.74 0.188 1.83 0.265 capns1 #2 1.432 0.182 1.91 0.228 calm1 #1 1.69 0.152 0.243 0.243 calm1 #2 1.14 0.127 0.0166 0.0166 calm2 #1 1.33 0.166 0.132 0.078 calm2 #2 1.22 0.183 0.0664 0.015 calm1 #1/calm2 #1 1.44 0.151 1.06 0.311 calm1 #2/calm2 #2 1.32 0.273 1.09 0.416 camk1 #1 1.09 0.299 0.203 0.145 camk1 #2 1.53 0.263 0.206 0.118 camk2a #1 0.97 0.264 0.252 0.284 camk2a #2 1.42 0.283 0.241 0.144 camk2b #1 1.39 0.211 0.311 0.188 camk2b #2 1.44 0.24 0.16 0.0981 camk1 #1/camk2a #1 1.89 0.174 1.02 0.204 camk1 #2 /camk2a #2 1.57 0.293 1.18 0.393 camk1 #1/camk2b #1 1.41 0.31 0.033 0.0242 camk1 #2/camk2b #2 1.29 0.283 0.17 0.0886 camk2a #1/camk2b 0.137 0.41 0.283 #1 1.33 camk2a #2/camk2b 0.152 0.22 0.144 #2 1.11 gnaq #1 1.2 0.273 0.751 0.22 gnaq #2 1.48 0.284 0.983 0.245 prkaca #1 1.223 0.236 0.203 0.306 prkacb #2 1.24 0.331 0.4 0.173 prkaca #1 1.05 0.364 0.133 0.0881 prkacb #2 1.94 0.236 0.13 0.0458

66 prkaca #1 /prkacb #1 1.77 0.164 0.116 0.0617 prkaca #2/prkacb #2 0.96 0.234 0.08 0.0404 prkca #1 1.24 0.236 0.0633 0.0633 prkca #2 1.05 0.231 0.113 0.094 prkcb #1 1.09 0.296 0.176 0.092 prkcb #2 1.34 0.369 0.08 0.0416 prkca #1/prkcb #1 1.69 0.253 0.943 0.233 prkca #2/prkcb #2 1.12 0.129 0.776 0.174 prkca #1/prkcg #1 1.53 0.244 0.18 0.126 prkca #2/prkcg #2 1.33 0.332 0.233 0.233 prkcb #1/prkcg #1 1.07 0.188 0.103 0.103 prkcb #2/prkcg #2 1.06 0.243 0.183 0.0652 plcb1 #1 1.4 0.238 0.11 0.104 plcb1 #2 1.6 0.385 0.176 0.0766 plcd1 #1 1.04 0.211 0.213 0.0551 plcd1 #2 1.13 0.283 0.118 0.0413 plcg1 #1 1.2 0.236 0.044 0.08 plcg1 #2 1.75 0.237 0.113 0.0133 plcb1 #1/plcg1 #1 1.26 0.235 0.733 0.375 plcb1 #2/plcg1 #2 1.14 0.384 0.693 0.196 plcb1 #1/ plcd1 #1 1.41 0.112 0.179 0.0551 plcb1 #2/plcd1 #2 1.67 0.295 0.143 0.0523 plcd1 #1/plcg1 #1 1.22 0.263 0.177 0.0989 plcd1 #2/plcg1 #2 1.31 0.254 0.136 0.0491 ! 67 Figure 7: siRNA screen identifies novel signaling components required for T. gondii-mediated cytolysis. (A, B)

B). B).

lls.(

mediated scoredcytolysis atare 60 -

T. gondii T.

Defects Defects in .

sis

tact vacuoles with >64 parasites) are compared at 44 and 60 hpi to identify induced cytoly -

Modelhost of screensiRNAmediatorsfor of parasite (A). ce new reinvade cells host exit to able parasites while parasites, >64 with vacuoles as persist exit to unable parasites hpi: (in vacuoles unruptured cells: in U2OS screen siRNA Primary SEM ± field per vacuoles unruptured mean are shown Data **p<0.01). (*p<0.05; release parasite in involvement gene host

Figure 7: siRNA screen identifies novel signaling components required for T. gondii-mediated cytolysis. (C-E)

correlation Pearson D). D).

knockdown shRNA stable within measurements doublings Parasite E). E). (

in resulted knockdown whose pairs gene and genes host dividual

in validates oligos multiple utilizing screen Secondary C). C). ( ( SEM. ± field per vacuoles mean unruptured are shown Data 60 hpi. by vacuoles unruptured of accumulation (r=0.91). oligos siRNA between relation strong indicates graph lines. cell

Figure 7: siRNA screen identifies novel signaling components required for T. gondii-mediated cytolysis. (F, G)

G). G).

. kdown cell lines at 44 and 60 hpi 60 and 44 at lines cell kdown mediated stable knockdown of target proteins, as compared to Scr oligo. ( - GFP knoc in vacuolesstable - Western blot analysis confirms shRNA F). F). ( of P30 images Representative

Figure 8. Design of siRNA screen to identify host genes required for T. gondii-mediated cytolysis.

For the primary screen, pooled siRNAs targeting a select set of 50 genes and 8 multiple gene families (three siRNAs/gene) were arrayed in collagen-coated 96-well plates (in quadruplicate), reverse-transfected into U2OS cells. Following synchronous T. gondii infection at 24 hours post-transfection, plates were fixed and imaged by fluorescence microscopy at 6, 24, 44 or 60 hpi. Image analysis was used to calculate parasite doublings as a measure of parasite replication and parasites per PV to assess progression of the entire intracellular parasite life cycle including cytolysis. Hits were selected based on persistence of large vacuoles at 60 hpi. The secondary screen included 19 individual genes and degenerate pairs of the multiple gene family hits, for a total of 17 multiple gene knockdowns using multiple oligos for validation. Hits from the secondary screen (individual genes or gene pairs) were tested in the final screen in 24-well plates. Images/western blots of stable shRNA knockdown cells were utilized in Figure 7F,G.

Figure 9: Conserved function of host proteins in P. falciparum- mediated cytolysis. (A, B)

cat cat β α / a images following immunodepletion immunodepletion following images a Representative Giems Representative

B).

1 results in persistence of schizont stage parasites by 60 hpi versus mock or PKA or mock versus 60 hpi by parasites stage schizont of persistence in results 1 - , and CaM β α / target protein elution from sepharose beads). (

– 1, PKC 1, β q, PLC q, α . Immunodepletion studies of soluble erythrocyte protein hits confirmed by western blot (M: Mock; ID: immunodepletion; IP: A) ( immunoprecipitation G host of hpi. 60 by reinvasion and exit in resulted which depletion,

Figure 9: Conserved function of host proteins in P. falciparum- mediated cytolysis. (C, D)

) parasites in depleted erythrocytes. Data shown are means stage stage (right - .01; *p<0.05). Data shown are means of at least three experiments ± SEM. . Ring parasitemia assessed 12 hours following needle shearing of infected, depleted D) stage (left) and stage (left) ring - Flow cytometric quantitation ofFlow quantitation cytometric schizont

C). ( of at least experiments ± three SEM ( (**p<0.01) **p<0 determined; Not (ND: indicated times the at erythrocytes 73

Figure 10. Depletion of PLCβ1 and calmodulin-1 from erythrocytes is sufficient to block P. falciparum-mediated cytolysis.

(A). Erythrocytes depleted of either PLCβ1, PLCγ, or both PLCβ1 and PLCγ were assayed for schizont persistence (left panel) or reinvasion of new rings (right panel) at 60 hpi. Though PLCβ1 depletion caused a significant persistence of schizont stage parasites and a reciprocal decrease in newly reinvaded rings (*p<0.05), indicating a block in cytolysis, PLCγ depletion did not alter normal life cycle progression. Dual depletion of both PLCβ1 and PLCγ did not significantly exaggerate the block in cytolysis from PLCβ1 alone, indicating that PLCγ does not have a significant role in this process. Data shown are means of at least 3 experiments +/- SEM; n=3. (B). Erythrocytes depleted of either calmodulin-1, calmodulin-2, or both calmodulin-1 and calmodulin-2 were assayed for schizont persistence (left panel) or reinvasion of new rings (right panel) at 60 hpi. Though calmodulin-1 depletion caused a significant persistence of schizont stage parasites and a reciprocal decrease in newly reinvaded rings, indicating a block in cytolysis (*p<0.05), calmodulin-2 depletion did not alter normal life cycle progression. Dual depletion of both calmodulin-1 and calmodulin-2 did not significantly exaggerate the block in cytolysis from calmodulin-1 alone, indicating that calmodulin-2 does not have a significant role in this process. Data shown are means of at least three experiments +/- SEM; n=3.

Figure 11: Host PKC is activated late in the parasite intracellular cycle and abrogates host adducin cytoskeletal association. (A-C)

Representative Representative FRET Representative control Representative B). B). A). A).

parasites. ( parasites.

P. falciparum P. depletion, depletion, 500 nM Gö6976. (

CKAR β α / - CFP/YFP CFP/YFP FRET fluorometry as a measurement of host PKC activity throughout C). C). nM nM PMA; negative controls: PKC

re mean fluorescence of at least three experiments ± SEM (**p<0.01). intracellular cycle. (

P. P. falciparum cycle. Data shown a shown Data cycle. . Assessment of host PKC activity through imaging and fluorometry of 3D7 P. falciparum falciparum P. C) 75 - A ( FRET images at 35 hpi. Positive control: 100 images through the the Figure 11: Host PKC is activated late in the parasite intracellular cycle and abrogates host adducin cytoskeletal association. (D-F)

D). D).

knockdown, 500 nM 500 knockdown,

β α / periments ± SEM (**p<0.01). SEM ± periments . CFP/YFP FRET fluorometry as a measurement of host infected U2OScells expressing cytoplasmic CKAR. ( - . (F) T. gondii T. intracellularcycle T. gondii gondii T. divisions. Positive control: 100 nM PMA; negative controls: PKC controls: negative PMA; nM 100 control: Positive divisions.

cycle. Data shown are mean fluorescence of at least three ex three least at of fluorescence mean are shown Data cycle. T. gondii gondii T. Representative FRET images through through the images FRET Representative

E. . Assessment of host PKC activity through imaging and fluorometry of F) - D

76 ( parasite at5 images FRET control Representative Gö6976. the throughout PKC activity

Figure 11: Host PKC is activated late in the parasite intracellular cycle and abrogates host adducin cytoskeletal association. (G, H)

treated erythrocytes (top) or -

2OS 2OS cytoskeletal fraction through

cycle in mock

P. P. falciparum

Western blot analysis of U

H).

stable knockdown cells (bottom). Loading control: actin. actin. control: Loading (bottom). cells knockdown stable -

β α /

(bottom). Loading control: actin. (

β α /

treated cells (top) or PKC -

cycle in mock in cycle

. Western blot analysis of erythrocyte cytoskeletal fraction through G) ( erythrocytes depleted of PKC gondii 77

Figure 11: Host PKC is activated late in the parasite intracellular cycle and abrogates host adducin cytoskeletal association. (I-L)

K). K). ). ( I

). I mutant adducin expression. - infection at 44 hpi, compared to

g host cell manipulation as in ). Data shown are means of at least 4 I T. T. gondii owing host cell manipulation as in as manipulation cell host owing vacuoles vacuoles at 60 hpi (followin T. T. gondii Parasite doublings throughout life cycle (foll cycle life throughout doublings Parasite

L). Representative Representative 60 hpi images of stable stable knockdown. Add1+: adducin overexpression; S716A/S726A: PKC phospho J). J).

prkca/prkcb or or

Western blot analysis of A549 cytoskeletal fractions upon adducin mutant expression, following I). ( capns1 Loading control: actin. ( Quantitation of unruptured vacuoles at 60 versus ( 44(**p<0.01). SEM ± experiments hpi (following host cell manipulation as in 78

Figure 12. Host Gαq and PLCβ1/γ1 are required upstream for host PKCα/β function in adducin loss prior to parasite-mediated cytolysis. (A, B)

. Data shown are

gondii T. and

viability upon mechanical mechanical upon viability

T. gondii T.

P. falciparum P. SEM.

ingblock of host cell cytolysis in hostcells uponGö6976

follow viability

T. gondii gondii T.

B). B).

SEM; n=3. n=3. ( SEM;

values and percent parasite reinvasion upon 500 nM treatment on on treatment nM 500 upon reinvasion parasite percent and values

mean are shown Data expression. adducing S716A/S726A or knockdown,

β α /

. PKC inhibitor IC (A) meansleast of three at experiments +/ treatment, PKC release at the timepoints shown following arrest at egress, of at least 3 experiments +/

Figure 12. Host Gαq and PLCβ1/γ1 are required upstream for host PKCα/β function in adducin loss prior to parasite-mediated cytolysis. (C-E)

(C). 3D7-CKAR parasites removed from their host cells via saponin treatment were assessed for CKAR fluorescence to account for digestive vacuole uptake. Representative 30-hour image (left) and population-based fluorometry throughout the life cycle (right) show digestive vacuole fluorescence, which is removed as background from all 3D7-CKAR measurements in Figure 12. Data shown are means of at least three experiments +/- SEM; n=4. (D). 3D7-CKAR fluorescence at 35 hpi following immunodepletion of key host signaling components. Representative images show that Gαq and PLCβ1 immunodepletion diminished CKAR signal in the host cell compartment, which was rescued by 100 nM PMA. Data shown are means of at least three experiments +/- SEM. (E). Erythrocyte CKAR FRET-based fluorescence of 3D7-CKAR parasites and life cycle progression as measured by quantitation of ring stage parasites at 60 hpi following depletion of Gαq or PLCβ1. Data shown are means of at least three experiments +/- SEM

Figure 12. Host Gαq and PLCβ1/γ1 are required upstream for host PKCα/β function in adducin loss prior to parasite-mediated cytolysis. (F,G)

tern blot

prkca/prkcb

dducin loss following following loss dducin following knockdown of

P. falciparum P.

ionsat 35hpi of

. Adducin immunofluorescence and western blot of U2OS G)

by immunofluorescence or western blot. PMA could not induce adducin loss following following loss adducin induce not could PMA blot. western or immunofluorescence by

plcb1/plcg1 or or

gnaq

. Adducin immunofluorescence and western blot of erythrocyte cytoskeletal fract F) ( wes or (left) immunofluorescence by β PLC or α q G of depletion following loss adducin rescues PMA components. signaling key (right). PMA could not induce adducin a rescues PMA loss following components. signaling key depletion of knockdown following of PKCdivisions /α β .parasite ( 5 at fractions cytoskeletal cell of knockdown knockdown. 81

Figure 12. Host Gαq and PLCβ1/γ1 are required upstream for host PKCα/β function in adducin loss prior to parasite-mediated cytolysis. (H,I)

based based -

. CKAR FRET I

components. signaling host SEM.

SEM.

knockdown diminished CKAR signal in the host cell compartment, compartment, cell host the in signal CKAR diminished knockdown

divisions following stable knockdown of key key of knockdown stable following divisions

T. gondii gondii T. plcb1/plcg1

gnaq infected U2OScells at5 parasite divisions. PMAaddition rescues PKCactivity following depletion - . Data shown are means of at least three experiments +/

T. gondii T. based fluorescence at 5 at fluorescence based -

plcb1/plcg1 or or

. CKAR FRET gnaq uorometry within within uorometry H) ( that show images Representative +/ experiments three least aremeansof at nM 100 byData rescuedPMA. shown was which fl of

Figure 13: Host signaling cascade enhances calpain-mediated cytoskeletal proteolysis just prior to parasite exit.

(A). DCG04-labeling of active calpain in P. falciparum-infected erythrocyte fractions at 50 hpi in cells depleted of Gαq, PLCβ1, PKCα/β, and CaM-1 by immunodepletion. Biotin blot confirms calpain activity upon labeling. Negative control: Calpain-1 depletion; positive control: PKAα/β depletion; loading control: actin. (B). α−Spectrin cleavage by western blot at 50 hpi in erythrocyte cytoskeletal fractions following depletion of signaling components. (C). α−Spectrin cleavage by western blot following in vitro incubation of U2OS cell cytoskeletal fractions with activated CaMKII prior to incubation with recombinant human calpain-1. (D). Host CAMKII/calmodulin-1 coimmunoprecipitation through the T. gondii life cycle and presence of p-CAMKII (the activated form) by western blot.

Figure 14: Host [Ca2+] increases throughout the last third of the intracellular cycle, in a TRPC6-dependent manner. (A) P. P. falciparum dextran dextran prior to challenge with synchronous - 2 - loaded with Fura

] as a function of 340nm/380nm emission ratio (right). Data shown are mean calcium measurements of 2+ . Representative images of erythrocytes A) ( [Ca (left). parasites SEM. ± 3 experiments least at 84

Figure 14: Host [Ca2+] increases throughout the last third of the intracellular cycle, in a TRPC6-dependent manner. (B)

] as a function of CFP/YFP 2+ infected U2OS cells expressing YC3.6 (left). [Ca measurementsof threeat experiments least ±SEM. -

2+ T. T. gondii s of Representative Representative image

B). ( Ca mean are shown Data (right). FRET

Figure 14: Host [Ca2+] increases throughout the last third of the intracellular cycle, in a TRPC6-dependent manner. (C,D)

(C) Representative images and quantitation (D) at 44 and 60 hpi of siRNA-mediated knockdown of cation channels prior to T. gondii infection. Positive control: Capns1; negative control: scr oligo. Data shown are mean unruptured vacuoles per field ± SEM (*p<0.05; **p<0.01).

Figure 14: Host [Ca2+] increases throughout the last third of the intracellular cycle, in a TRPC6-dependent manner. (E)

(E). Representative images (left) and [Ca2+] measurements as a function of CFP/YFP FRET (right) at 5 T. gondii divisions (~35 hpi) following siRNA-mediated knockdown of cation channels in U2OS cells expressing YC3.6. Data shown are mean Ca2+ measurements ± SEM.

Figure 15. PKC activity, as mediated by Gαq and PLC, is required for Ca2+ influx at the end of the parasite life cycle. (A, B)

A). Giemsa images at specified timepoints following treatment beginning at 30 hpi with 500 nM cation channel inhibitors GsTMx-4, 2-APB, SKF-63563, or Pyr3. (B). Population level measurement of schizont persistence (left) and new ring formation (right) following inhibition of cation channel inhibitors at 30 hpi. Data shown are means of at least three experiments +/- SEM; *p<0.05; **p<0.01.

Figure 15. PKC activity, as mediated by Gαq and PLC, is required for Ca2+ influx at the end of the parasite life cycle. (C)

(C). Representative FRET images and population level fluorometry at 5 T. gondii parasite divisions of U2OS cells expressing YC3.6 following knockdown of gnaq, plcb1/plcg1, or prkca/prkcb show that [Ca2+] is limited upon knockdown of these signaling members. 100 nM PMA treatment rescues Ca2+ influx following gnaq or plcb1/plcg1 knockdown, though it cannot be rescued following prkca/prkcb knockdown. Data shown are means of at least three experiments +/- SEM.

Figure 15. PKC activity, as mediated by Gαq and PLC, is required for Ca2+ influx at the end of the parasite life cycle. (D)

(D). Representative 45 hpi images and population-level Ca2+ measurements of P. falciparum-infected 2+ erythrocytes loaded with Fura-2-dextran, and immunodepleted of Gαq, PLCβ1, or PKCα/β. Ca influx was limited in cells depleted of these effectors, but could be rescued upon PMA treatment following Gαq or PLCβ1 depletion. The mechanosensitive channel inhibitor GsTMx-4 abrogated PMA-induced Ca2+ influx, indicating that is influx is cation channel-dependent. Data shown are means of at least three experiments +/- SEM.

Figure 16

Figure 16: Mammalian PKC inhibitors show antiparasitic activity in vivo. (A). P. yoelii parasitemia quantified via Giemsa tail vein blood smear upon intravenous injection of 10 mg/kg Gö6976 once daily for 4 days. (n=10; ***p<0.001). (B). T. gondii parasite burden in cells from the spleen (top) and peritoneal cavity (PECS; bottom) upon intraperitoneal injection of 10 mg/kg Gö6976, quantified by flow cytometry. Plots shown are gated on single, live cells by FSC and SSC. (n=15 for all conditions; ***p<0.0001). (C). P. berghei ANKA parasitemia quantified via Giemsa tail vein blood smear upon 50 mg/kg Sotrastaurin treatment via gastric gavage once daily for 4 days. (n=7 for all conditions; ***p<0.0001). Shown are mean parasitemias ± SEM. (D). Survival curves following sotrastaurin treatment using the standard Mantel-Cox log rank test (**p=0.0029).

Figure 17: Parasite TCA cycle intermediates initiate the host cytolytic network. (A, B).

Immunofluorescence of OXGR1 and SUCNR1 within P. falciparum-infected erythrocytes (A) and T. gondii– infected U2OS cells (B). Western blots are included as a control. (A: Red- OXGR1 (top panel)/SUCNR1 (bottom panel), Green- His-GFP (PV); (B): Red– OXGR1 (top panel)/SUCNR1 (bottom panel), Green- P30- GFP (PV).

Figure 17: Parasite TCA cycle intermediates initiate the host cytolytic network. (C,D)

(C). Western blot confirmation of siRNA-mediated knockdown of host GPCRs and the positive control CAPNS1. Representative images of P30-GFP-infected U2OS knockdown cells at 44 and 60 hpi. (D). Quantitation of enlarged vacuoles through 60 hpi. Data shown are means of at least three experiments +/- SEM.

Figure 17: Parasite TCA cycle intermediates initiate the host cytolytic network. (E-G)

(E,F). Host OXGR1 and SUCNR1 activity following T. gondii infection via measurement of Fluo-8 fluorescence. Representative images (E) and Fluo-8 fluorescence (F) indicate host GPCR activity throughout the parasite life cycle. Data shown are mean fluorescence +/- SEM. (G) Unruptured vacuoles measured at 48 hpi following T. gondii infection of cells used in (E, F).

Figure 17: Parasite TCA cycle intermediates initiate the host cytolytic network. (H, I)

(H). Time until T. gondii parasite exit upon treatment with 50 µM exogenous aKG/succinate or 50 µM lactate, versus 10 µM A23187 (positive control for egress induction). Data shown are mean time of at least 3 experiments +/- SEM. (I). Representative Giemsa images following a 12-hour treatment with 50 µM exogenous aKG/succinate, lactate, or DMSO control.

Figure 18: Model of host GPCR-mediated cytolysis.

Parasite ligands, perhaps upregulation of parasite TCA cycle intermediates during periods of high parasite replication, activate GPCRs on the PV membrane and initiate a signaling cascade through host Gαq. PKC- mediated phosphorylation liberates adducin from the host cell cytoskeleton, activating the mechanosensitive plasma membrane cation channel TRPC6. CaMKII-activation following Ca2+ influx mediates phosphorylation of cytoskeletal substrates, which, upon rapid influx of Ca2+ from the extracellular media, are proteolyzed by calpain to allow for cytoskeletal dissolution and parasite release. 96

VI. Human Platelet Factor 4 and synthetic mimics: Harnessing host defense mechanisms for parasite death by ulcer The pathogenesis of malaria is modulated both positively and negatively by many human cell types in blood including monocytes, neutrophils and platelets (de Mast et al.,

2007; Gimenez et al., 2003; Goodier et al., 1995; Krupka et al., 2012; Mohanty et al.,

1997; Pain et al., 2001). While platelets have been long known to interact cooperatively with erythrocytes in thrombogenesis, both generally and in the specific case of sickle cell disease (Rocca and FitzGerald, 1997), only recently have platelets been shown to bind infected erythrocytes and kill intracellular malaria parasites (McMorran et al., 2009).

Previous studies have suggested that thrombocytopenia is a poor prognostic marker in malaria (Moerman et al., 2003) and is significantly associated with cerebral malaria (Cox and McConkey, 2010; Wassmer et al., 2008). We were intrigued by the idea that these host cells found in the bloodstream are known to secrete proteins with host defense peptide activity (HDPs) (McMorran et al., 2009; Tang et al., 2002), and the possibility that these proteins could be responsible for controlling malaria parasite proliferation in the blood stage.

HDPs play a central role in the innate immune system (Finlay and Hancock, 2004;

Hancock and Lehrer, 1998; Tossi et al., 2000; Zasloff, 2002). Over 700 HDPs have been identified, varying in length, sequence, and tertiary structure. There are two classes of

HDPs comprising ribosomally and nonribosomally synthesized peptides: α-helical (i.e. magainin and cecropin) and disulfide-rich β-sheets (i.e. bactenecin and defensin), although other tertiary structures also exist. Importantly, HDPs display broad-spectrum action against bacteria, fungi, protozoa, and viruses, which has promoted their use as new leads for developing antibiotics (Zasloff, 2002). HDPs share common features

97 including amphipathic topologies in which the cationic and hydrophobic side chains segregate onto distinctly opposing regions or faces of the overall folded conformation.

Furthermore, numerous studies with linear and cyclic peptides have strongly supported the hypothesis that the physicochemical properties of HDPs, rather than any specific sequence or structure, are responsible for their activities. HDPs are thought to bind the membrane surface in a non-cooperative fashion and then aggregate once a threshold concentration is reached causing membrane permeabilization (Christensen et al., 1988;

Ludtke et al., 1995; Matsuzaki et al., 1995, 1996; Pouny et al., 1992). It is generally believed that amphipathic topology is essential for insertion into and disruption of membranes leading to pathogen death (Zasloff, 2002). HDPs have remained an effective weapon against pathogen infection over time indicating that their mechanism of action thwarts pathogen responses, which normally lead to resistance against toxic substances. This premise is supported by data showing that no appreciable resistance to the action of the HDPs occurs after multiple serial passages of bacteria in the presence of sub-lethal concentrations of the peptides (Ge et al., 1999; Mosca et al.,

2000; Tew et al., 2002). From this background, we hypothesized that naturally occurring bloodstream HDPs especially from platelets could be responsible for killing of P. falciparum early in erythrocytic infection, aside from cerebral malaria symptomology.

Identification of human platelet factor 4 (hPF4) as an antiparasitic HDP that kills P. falciparum via lysis of the parasite DV

A screen of HDPs normally found in the bloodstream, secreted by a variety of cells including platelets, neutrophils, or lymphocytes, was performed to assess their antiparasitic activity (Figure 19A). This screen revealed that several proteins killed P.

98 falciparum in vitro without affecting the host erythrocyte. Most notably, hPF4 showed

high potency against P. falciparum with an IC50 of 4.2 µM and no significant hemolysis.

Considering local concentrations of hPF4 have been reported to reach at least 280 µM surrounding activated platelets (Kowalska et al., 2010), the antiparasitic IC50 of hPF4 has in vivo relevance. We thus focused our efforts on studying PF4 to determine the biological relevance of its potential antimalarial properties. Platelets harvested from wild- type (WT), mouse PF4 (mPF4) knock-out (mPF4 KO), or overexpressing hPF4 (hPF4+) mice were tested against P. falciparum in culture with recombinant mPF4 and hPF4 as controls (Figure 19B). Pre-activated WT and hPF4+ mouse platelets were able to kill P. falciparum in culture, while littermate PF4 KO platelets showed no killing capacity. Thus it appears that PF4 is the major antimalarial component of activated platelets.

As membrane perturbation is an established mechanism of action for HDPs

(Burkhard, 2009; Lehrer et al., 1989; Matsuzaki et al., 1997; Westerhoff et al., 1989;

Zasloff, 2002), we examined the integrity of potential target parasite membranes upon hPF4 treatment. Since hPF4 had little hemolytic capability we reasoned the erythrocyte integrity was not significantly compromised. Parasite plasma membrane (PPM) potential assays showed no discernible loss of potential following 4 hours of hPF4 exposure

(Figure 19C). Next the integrity of established intracellular organelle targets, including the mitochondria and lysosome-like DV was assessed post hPF4 treatment. Analysis of membrane potential revealed no perturbation of the mitochondria, however a significant loss of proton potential was observed in the DV within minutes of treatment (Figure

19C). To further investigate this novel perturbation to the DV, transgenic parasites expressing green fluorescent protein (GFP)-tagged plasmepsin II in the DV (Klemba et al., 2004) were treated with hPF4 and followed by fluorescence microscopy over a 10- 99 minute time course. Parasites were assayed for both hPF4 localization using immunofluorescence analysis (IFA) and direct GFP signal (integrity of the DV; Figure

19D). Within 1 minute of hPF4 incubation, hPF4 staining could be detected in the cytoplasm of only infected erythrocytes. hPF4 continued to accumulate over the next several minutes, wherein it entered the parasite cytoplasm and colocalized with the parasite DV. Next, severe perturbation of the DV allowed GFP to diffuse throughout the parasite cytoplasm (Figure 19D, Figure 20A,B). Western blot analysis of hPF4 confirmed the IFA results showing entry into erythrocytes by 1 minute followed by accumulation into the parasite within 5 minutes (Figure 19E). To assess the loss of DV integrity on the population level, we utilized ImageStream flow cytometry to first capture images of individual parasite-infected cells, then quantitatively differentiate based on DV fluorescent surface area (SA) intact DVs (<0.2 µm2) from lysed DVs (>2 µm2; Figure

19F, Figure 20C). hPF4 treatment caused dose-dependent DV lysis, with nearly complete DV lysis on a population level upon exposure to 10 µM hPF4. The loss of DV integrity was confirmed at high resolution using transmission electron microscopy imaging to compare untreated and hPF4-treated parasites revealing the complete loss of

DV integrity in the hPF4-treated parasites versus mock-treated parasites with a clearly delineated DV membrane (white arrowheads; Figure 19G). In addition to the loss of DV organellar structure, hemozoin crystals were seen dispersed throughout the parasite (red arrowheads). Thus, DV lysis appears to be the primary and novel mechanism of hPF4 action against parasites.

In contrast, the only other human HDP tested against P. falciparum is NK-2, secreted by lymphocytes, which was shown to kill P. falciparum by direct perturbation of the

100 infected erythrocyte plasma membrane (Gelhaus et al., 2008). Dermaseptins, HDPs from the skin of hylid frogs, were demonstrated to kill parasites by perturbation of the infected erythrocyte plasma membrane or the parasite plasma membrane (Dagan et al.,

2002; Ghosh et al., 1997). However, the high hemolytic potential of dermaseptins and the fact they are not of human origin makes the therapeutic relevance of these findings unclear. Though the mode of entry into cells is unclear, given that there are 4 membrane systems that must be bypassed prior to selective interaction with the DV membrane, we provide evidence that hPF4 enters only infected erythrocytes and that this entry process can be blocked by protamine sulfate (Figure 20D). This indicates the importance of initial electrostatic interactions with the infected host cell membrane or membrane proteins. Stage specificity studies indicate that hPF4 is most potent during parasite stages that heavily rely on digestive vacuole function (Figure 20E), perhaps indicating that hPF4 is incorporated into the parasite body during hemoglobin uptake.

In order to obtain proof of concept that HDPs kill parasites in blood stage malaria infection, we tested PF4 in a non-experimental cerebral malaria (non-ECM) murine malaria model using the 4-day Peters suppression test (Knight and Peters, 1980). Mice infected with P. yoelii 17XNL were administered 10 mg/kg mPF4 intravenously on days

1-3, and parasitemias were assessed on day 4. mPF4 treatment significantly reduced parasitemia by 4-fold (Figure 20F).

Synthetic small molecule HDP mimics are potent against P. falciparum and have a mechanism similar to hPF4

Given their broad activity, HDPs appear to be ideal therapeutic agents, though significant pharmaceutical issues have severely hampered clinical progress, including

101 poor tissue distribution, systemic toxicity, and difficulty and expense of manufacturing

(Easton et al., 2009). In addition, PF4 has been reported to worsen experimental cerebral pathology in mice, presumably mediated through the chemokine domain

(Srivastava et al., 2008). Thus, we reasoned that synthetic small molecules capable of adopting amphipathic secondary structures analogous to an HDP could potentially reproduce the potent, selective antiparasitic activity of PF4, while improving tissue distribution and decreasing complications arising from chemokine signaling. In addition, these small molecule HDP mimics (smHDPs) are significantly less expensive to produce

(an example scaffold is shown in Figure 21A). In order to exploit the DV lysis mechanism of PF4 in a drug discovery effort, a library of ~1000 non-peptidic smHDPs

(Mensa et al., 2011; Scott et al., 2008) was tested against P. falciparum. 132 compounds were determined to be initial “hits” (14.3% hit rate), using a cutoff of 80% parasite death at 1 µM (Z’ = 0.720; Figure 21B). 74 compounds were selected for follow-up IC50 determination (Table 6). Based on these hits, additional compounds were synthesized and 7 structurally diverse compounds were chosen as “leads” based on high potency against P. falciparum, low hemolytic potential, and low mammalian cell cytotoxicity. These lead compounds also showed similar potency across a panel of chloroquine-sensitive (3D7, HB3) and chloroquine-resistant (Dd2, 7G8, K1) parasite lines, both lab-adapted (Table 7) and short-term adapted field isolates (Table 8).

Several of the lead compounds displayed little activity against Gram-positive and Gram- negative bacteria, as well as mammalian cells, indicating that a high level of specificity can be built into these non-peptidic scaffolds that can distinguish between prokaryotic and eukaryotic pathogens. Although all lead compounds showed a similar mechanism of

102 action as PF4, based on acute animal tolerability studies, PMX1207 and PMX207 were selected for more rigorous mechanistic analysis.

Treatment of P. falciparum parasites with either PMX1207 or PMX207 disrupted DV membrane integrity within 15 minutes of treatment but did not disturb the PPM or mitochondrial potential, even after 4-hour treatment (Figure 22A). In addition, treatment of the transgenic PMII-GFP parasites with PMX1207 and PMX207 resulted in rapid diffusion of GFP through the parasite cytoplasm indicating a catastrophic loss of DV membrane integrity in a manner similar to that observed for hPF4 (Figure 22B).

ImageStream flow cytometry analysis indicated a dose-dependent effect on DV lysis, with nearly complete DV lysis within the treated population upon treatment with 1 µM of

PMX207 or PMX1207 (Figure 22C). Similarly, 7 additional smHDPs showed rapid lysis of the DV upon treatment (Figure 23A).

We further investigated the intracellular localization and distribution of the smHDPs using PMX496, a potent smHDP with intrinsic fluorescence. PMX496 appeared to accumulate in the parasite body within 30 seconds, then concentrated in the DV at 1 minute, prior to dissipation throughout the parasite body, indicating a loss of DV membrane integrity (Figure 23B). Since cationic amphipathicity is predicted to be necessary for smHDP antiparasitic action, an uncharged isosteric analog of PMX496 was synthesized (PMX1269, IC50 > 2.5 µM). As expected, treatment with the uncharged analog did not result in DV lysis. High-resolution transmission electron microscopy and

3D tomograms revealed a complete loss of integrity of the DV after treatment with either

PMX1207 or PMX207 (Figure 22E). In addition to free hemozoin crystals (red arrowheads), images showed an accumulation of undigested hemoglobin-containing

103 vesicles (yellow arrowheads; Figure 22E, Figure 23E). Mock-treated parasites showed clear delineating DV membranes (white arrowheads). DV lysis and killing was also evident in Texas Red Dextran-loaded erythrocytes infected with either trophozoite-stage parasites (Figure 23C) or gametocytes (Figure 23D). Although we cannot discern whether gametocytes are dying from this data, loss of DV integrity would likely result in death and this possibility indicates that smHDPs may have transmission-blocking potential. smHDP activity reduces parasitemia in a murine malaria model

In order to further explore the antimalarial properties of lead smHDPs, PMX207 and

PMX1207 were assessed for their ability to limit parasite growth in vivo. PMX207 and

PMX1207 antiparasitic activity was assessed in a non-ECM murine malaria model using the 4-day Peters suppression test (Figure 22E). Mice were inoculated intravenously on day 1 with 2 x 105 P. yoelii XNL-infected erythrocytes and then treated with 10 or 20 mg/kg of PMX207, 5 mg/kg PMX1207, or vehicle alone for 3 days and parasitemias were monitored on day 4. PMX207 showed a dose-dependent decrease in parasite growth, with 20 mg/kg, almost completely eliminating parasitemia (Figure 22F,G). PMX1207 showed a significant decrease in parasite growth, suppressing parasitemia 5-fold over vehicle-treated controls. These data provide in vivo validation that smHDP compounds show promise as a new class of antimalarials.

Discussion

HDPs are extremely diverse not only in structure, but in cellular targets and mode of action. HDPs have been shown to kill pathogens via a variety of mechanisms: simply via perturbation of the pathogen plasma membrane (Chao and Raines, 2011; Duchardt et

104 al., 2007; Haines et al., 2009; Mochon and Liu, 2008) or intracellular membranes such as mitochondria (Slaninova et al., 2012), or through immune modulation (D'Este et al.,

2012; Repeke et al., 2011; Williams et al., 2012; Wuerth and Hancock, 2011). However, there is a distinct lack of examples that involve perturbation of pathogen endosomes/lysosomes, aside from the killing of the distantly related protozoan parasite,

T. brucei brucei, in an HDP-like manner by a human protein, trypanosome lytic factor

(TLF). TLF has been shown to cause lysis of parasite endosomes/lysosomes conferring a species barrier and protection for humans against infection (Harrington et al., 2009;

Rifkin, 1978; Samanovic et al., 2009; Smith et al., 1995).

Finally, we present a model for the mechanism of action for these HDPs or small molecule mimics, which begins upon uptake of these molecules into the host erythrocyte cytoplasm, possibly through a parasite-mediated mechanism (Figure 24A). Once inside the host cell cytoplasm, these HDP molecules are targeted to the DV along with hemoglobin through the cytostome (Goldberg, 1993) and carried to the DV via hemoglobin-containing vesicles. Once inside the DV, the HDP or smHDP is released along with the hemoglobin cargo, accumulates and binds to the membrane directly. At a critical concentration, these HDPs disrupt the integrity of the DV membrane and release the contents, which include numerous proteases and hydrolases, into the cytoplasm of the parasite (Figure 24B). Destruction of proteins and organelles leads to rapid parasite death (Figure 24C).

105

Table 6

Compound 3D7 IC50 (nM) Dd2 IC50 (nM) Compound 3D7 IC50 (nM) Dd2 IC50 (nM) Chloroquine 8.7 24.6 614 864 966 Artesunate 19.4 12.1 616 149 97 004 563 740 617 110 172 015 438 601 633 272 457 018 668 611 647 160 300 033 306 419 651 148 267 048 295 484 661 175 200 053 740 1154 730 1150 975 056 653 993 731 1164 964 058 1233 1219 734 610 980 062 413 426 781 147 207 073 204 274 800 610 431 092 1024 713 819 724 951 112 688 740 832 358 368 143 870 688 835 220 300 146 322 244 847 870 1076 152 221 170 848 625 709 158 683 827 868 881 886 160 151 168 893 200 301 192 1198 1010 1007 1283 2747 201 125 85 1011 880 795 207 153 134 1012 656 555 217 569 591 1032 1197 798 224 893 524 1045 706 398 235 146 108 1064 1098 1132 256 1064 344 1065 680 664 270 732 803 1068 644 682 459 1365 1632 1070 650 724 462 1283 2120 1076 N/A N/A 474 991 1226 1086 812 1054 484 655 1095 1088 734 602 485 841 1006 1095 1141 786 493 904 1183 1096 58 90 496 160 170 1099 1163 667 501 593 393 1101 672 656 504 160 118 1107 1102 997 610 N/A N/A 1115 696 829 611 60 49 1120 308 331

1154 530 N/T 1284 160 135 1158 563 N/T 1287 2819 N/T 1159 493 N/T 1301 226 N/T 1164 N/A N/T 1302 160 132

106

1165 161 N/T 1305 N/A N/T 1170 81 N/T 1306 1299 N/T 1179 1125 N/T 1307 1256 N/T 1185 15.5 N/T 1308 629 N/T 1199 2932 N/T 1310 2203 N/T 1200 2681 N/T 1324 602 N/T 1201 3376 N/T 1325 112 N/T 1206 3284 N/T 1338 N/A N/T 1207 110 75 1364 N/A N/T 1209 1503 N/T 1374 641 N/T 1210 658 N/T 1375 88.3 N/T 1211 976 N/T 1382 126 N/T 1213 N/A N/T 1387 N/A N/T 1219 3050 N/T 1398 131 N/T 1220 N/A N/T 1399 N/A N/T 1221 N/A N/T 1400 473 N/T 1222 3603 N/T 1403 116 N/T 1223 3079 N/T 1407 795 N/T 1224 297 N/T 1408 77.1 N/T 1226 2523 N/T 1410 N/A N/T 1229 202 N/T 1411 N/A N/T 1237 255 N/T 1412 N/A N/T 1249 1716 N/T 1413 N/A N/T 1258 247 N/T 1414 N/A N/T 1260 N/A N/T 1415 N/A N/T 1267 N/A N/T 1416 N/A N/T 1268 3480 N/T 1417 N/A N/T 1269 3029 N/T 1418 187 N/T 1271 2318 N/T 1422 738 N/T 1283 2445 N/T N/A = Not Active N/T = Not Tested

Table 6. IC50 values determined in 3D7 (CQ-sensitive) and Dd2 (CQ-resistant) P. falciparum for 74 hits from smHDP screen and derivative compounds not included in original screen. Parasites were treated in triplicate, n = 2.

107

Table 7

Table 7. smHDPs show potency across several chloroquine-sensitive and chloroquine-resistant P. falciparum strains, with little cytotoxicity against mammalian cells. Lead smHDPs were screened against a panel of chloroquine-sensitive and chloroquine-resistant P. falciparum lines. Chloroquine was used as a positive control for parasite death. IC50s are as mean ± SEM, n>3 for each compound. Cytotoxicity (EC50) determined against mouse 3T3 fibroblasts and human transformed liver HepG2 cells using an MTS viability assay.

Table 8

Table 8. Lead smHDPs show potency across several chloroquine-sensitive and chloroquine- resistant P. falciparum field isolates. Lead smHDPs were screened against a panel of chloroquine- sensitive and chloroquine-resistant P. falciparum lines. Chloroquine was used as a positive control for parasite death. IC50s are as mean ± SEM, n>3 for each compound. Cytotoxicity (EC50) determined against mouse 3T3 fibroblasts and human transformed liver HepG2 cells using an MTS viability assay.

108

Figure 19. hPF4 acts as a HDP against P. falciparum via lysis of the parasite DV. (A,B)

shown shown ometry ometry

B); B); Human Neutrophil

B (FP -

platelets from either littermate WT, WT, PF4 littermate either from platelets

A); A); Fibrinopeptide

A (FP -

killing and hemolysis at 15 M, µ including Regulated upon Activation,

te survival was normalized to an untreated control. Artesunate (500 nM)

arum

were incubated for 24 hours with 2.5 x for 10hours2.5 were x 24 incubated with

37). Parasi -

P. falcip

A content and normalized an to control. hPF4Recombinant untreated and mPF4 (10 µ M)

2); Cathelicidin (LL2); Cathelicidin infected erythrocytes -

1; 1; HNP - ) Parasite

cell Expressed, and Secreted (RANTES); hPF4; Fibrinopeptide -

) Screen of human HDPs found in blood for A ( Normal T Peptides 1 and 2 (HNP and the HDP melittin (15 µ M) were used as positive controls for parasite death without and with hemolysis, respectively. Data SEM. ( ± means are KO, or hPF4+ mice in the presence or absence of 5 µ M AYP to induce platelet activation. Parasite survival assayed by flow cyt SYTOX using as of a green measurement DN SEM.means± Data shown are controls. positiveas were used

109

Figure 19. hPF4 acts as a HDP against P. falciparum via lysis of the parasite DV. (C-E)

(C) Parasite-infected erythrocytes pre-incubated with rhodamine 123 (1 µM for parasite plasma membrane potential; 0.2 µM for mitochondrial potential) or 10 nM LysoTracker Red and then treated over a 4-hr time course with 10 µM PF4 or a 10 µM mixture of ionophores Monensin and Nigericin (Mon/Nig). Length bar is 10 µm in each figure. (D) hPF4 accumulates in the infected erythrocyte cytoplasm prior to lysis of the DV by immunofluorescence. Erythrocytes infected with parasites expressing plasmepsin-II-GFP (PM-II-GFP) were treated with 1 µM hPF4 over a 10 min time course. Within 1 minute of PF4 incubation, PF4 staining is seen in the cytoplasm of infected erythrocytes until entering and inducing lysis of the parasite DV, as shown by GFP signal diffusion through the entire parasite cytoplasm. (Green: PM-II-GFP [DV]; red: hPF4 IFA; blue: parasite nuclei.) (E) Western blot analysis of erythrocyte fractions or parasite lysates shows early accumulation of hPF4 in infected erythrocytes, followed by persistence in the parasite.

110

Figure 19. hPF4 acts as a HDP against P. falciparum via lysis of the parasite DV. (F, G)

(F) ImageStream flow cytometry shows a dose-dependent increase in PM-II-GFP parasites with a fluorescence surface area of >2 µm2, an indication of DV lysis, upon treatment with hPF4. Intact DVs have fluorescent surface areas <0.2 µm2. Data shown are means ± SEM. (*p<0.05, **p<0.01) (G) TEM images reveal dissolution of the DV membrane upon treatment with 10 µM PF4, as well as dispersal of hemozoin fragments throughout the cytoplasm (red arrowheads). Mock-treated controls showed complete integrity of the DV membrane (white arrowheads), encapsulating all hemozoin crystals. (N: parasite nucleus).

111

Figure 20. hPF4 causes dose-dependent DV lysis and reduces parasitemia in a mouse malaria model. (A-C)

(A) Trophozoite-infected Texas Red Dextran-loaded erythrocytes show dispersal of fluorescence into the parasite cytoplasm (indicating DV lysis) following 5 min of treatment with hPF4. (B) Gametocyte-infected Texas Red Dextran-loaded cells show dispersal of fluorescence into the parasite cytoplasm (indicating DV lysis) upon 5 min of treatment with hPF4. (C) Representative flow plots from mock-treated parasites expressing PM-II-GFP or parasites treated with 1 µM hPF4, 3 µM hPF4, or 10 µM hPF4. Synchronous PM-II parasites were used to examine DV integrity upon treatment. DV lysis is defined as area of GFP signal greater than 2 µm2 (red line is minimum area), as GFP signal would disperse into the entire parasite cytoplasm. Intact DVs show compact area (less than 0.2 µm2; blue line is maximum area).

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Figure 20. hPF4 causes dose-dependent DV lysis and reduces parasitemia in a mouse malaria model. (D, E)

(D) PM-II-GFP parasite-infected erythrocytes were pretreated with 5 mM protamine sulfate or mock (DMSO) or 30 minutes, then treated with 10 µM hPF4 for 10 minutes prior to fixation and permeabilization. hPF4 immunofluorescence (red) shows no appreciable signal in the protamine sulfate pretreated erythrocytes, while mock-pretreated erythrocytes show hPF4 accumulation within the erythrocyte. (red: hPF4 IFA; green: PM-II-GFP; blue: parasite nuclei). (E) Stage-specific killing of PF4 shows increased efficacy during the stages that rely heavily on DV function (late ring, trophozite, schizont, gametocyte). Shown are means ± SEM; n=3.

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Figure 20. hPF4 causes dose-dependent DV lysis and reduces parasitemia in a mouse malaria model. (F)

(F) Mice infected with P. yoelii 17XNL show decreased parasitemia on day 4 when treated with 10 mg/kg mPF4 (n=3) compared to controls (n=5, p < 0.001). Shown are mean parasitemias ± SEM.

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Figure 21. Screen of smHDPs reveals potent inhibitors of P. falciparum growth.

(A) Conceptual design of smHDPs from HDPs. Top: Amphipathic structure of magainin 2; cationic groups in blue, nonpolar groups in green, peptide backbone in yellow. Bottom: de novo designed smHDPs capture the facially amphipathic architecture and critical physicochemical properties needed to establish robust antimicrobial activity. (B) Screen of 920 smHDPs at 500 nM against P. falciparum. Parasite death was normalized to a chloroquine control (14.3% hit rate, Z’ = 0.720).

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Figure 22. smHDP leads kill P. falciparum via parasite DV lysis and decrease parasitemia in a murine malaria model.

(A) Parasite plasma membrane and mitochondrial potential were examined during treatment with PMX207 or PMX1207, with no discernible loss of fluorescence, unlike the positive controls. Compromise of DV integrity was monitored with LysoTracker Red, wherein loss of DV fluorescence was seen within 15 min of smHDP treatment, though no loss of parasite plasma membrane potential occurs. (B) Loss of DV integrity 116 upon smHDP treatment of PM-II-GFP P. falciparum parasites (Green: PMII-GFP [DV]; blue: parasite nuclei.) (C) ImageStream flow cytometry shows a dose-dependent increase in PM-II-GFP parasites showing a fluorescence surface area of >2 µm2, an indication of DV lysis, upon treatment with PMX207 or PMX1207. Intact DVs have fluorescent surface areas <0.2 µm2. Data shown are means ± SEM. (*p<0.05, **p<0.01) (D) TEM images reveal complete dissolution of the DV membrane upon PMX207 or PMX1207 treatment, with dispersal of hemozoin crystals (red arrowheads) throughout the cytoplasm and an accumulation of undigested hemoglobin-containing vesicles (yellow arrowheads). Mock-treated parasites retained a clear delineating DV membrane (white arrowheads), encapsulating all hemozoin crystals. (N: parasite nucleus). (E) Parasitemias of Swiss Webster mice infected with P. yoelii 17XNL parasitized erythrocytes via i.v. injection and treated with vehicle (n=5); 5 mg/kg PMX1207 (n=7); or 20 mg/kg of PMX207 (n=5). Parasitemia was assessed on days 4 and 6. Shown are the means ± SEM. (*p<0.05; ***p<0.001) (F) Parasitemias of C57Bl/6 mice infected with P. berghei ANKA parasitized erythrocytes via i.v. injection and treated with vehicle (n=10); 5 mg/kg PMX1207 (n=5); or 20 mg/kg of PMX207 (n=5). Parasitemia was assessed on days 5 and 7. Shown are the means ± SEM. (*p<0.05; ***p<0.001). (G) Survival curves for P. berghei ANKA infected mice. p = 0.14 in the log-rank Mantel-Cox test.

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Figure 23. smHDPs require a facial charge density to lyse the parasite DV.

(A) Visualization of DV lysis by smHDPs by fluorescence microscopy. PM-II-GFP P. falciparum parasites were treated with compounds PMX504, PMX611, PMX647, PMX835, PMX496, or PMX1269 (250 nM) over a 5-min time course and imaged via fluorescent microscopy. In untreated parasites, the GFP signal is an isolated structure, indicating an intact DV. The GFP signal after smHDP treatment spans the entire parasite body, indicating loss of digestive vacuole integrity. The uncharged analogue of PMX496, PMX1269, did not cause DV lysis. (Green: PM-II-GFP; blue: parasite nuclei.) (B) Fluorescent microscopy of the intrinsically fluorescent PMX496 shows accumulation within the parasite cytoplasm and DV prior to DV lysis. (C,D) Visualization of DV lysis by PMX1207 with Texas Red Dextran-loaded erythrocytes. Trophozoite- (C) or Gametocyte-infected (D) Texas Red Dextran-loaded cells show dispersal of fluorescence into the parasite cytoplasm following treatment with PMX1207, indicating loss of DV integrity. (E) Number of hemoglobin- containing vesicles in electron micrographs of parasites treated with 10 µM hPF4, 500 nM PMX207, or 118

500nM PMX1207 was counted and compared to mock (DMSO)-treated controls. Data shown is mean ± standard deviation of 50 parasites (*p<0.05).

Figure 24

Figure 24. Model of PF4 and smHDP uptake. Proposed mechanism of parasite uptake of PF4 and smHDPs. (A) Molecules reach the host erythrocyte cytoplasm, possibly via either an active mechanism (Route 1) or a passive mechanism (Route 2) initiated by the parasite during hemoglobin uptake through the cytostome (c) and packaging into double membranous hemoglobin-containing vesicles (HV). HVs are then targeted to the DV where the contents are released into the DV lumen. (B) As the parasite continues to take up host cell cytoplasm, the HDPs or derivatives eventually reach a critical concentration within the DV, bind to the inner leaflet of the DV membrane, and disrupt its integrity, allowing the contents to enter the cytoplasm of the parasite. (C) Rapid destruction of proteins and organelles cause parasite death.

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VII. CONCLUSIONS AND FUTURE DIRECTIONS

Through focused characterization of host-parasite interactions that could be exploited to strengthen development of novel antiparasitics, we hoped to achieve a paradigm shift to establish that multiple host proteins/pathways are essential for apicomplexan parasite growth, and may be exploited to generate novel drug classes with limited propensity for resistance. In addition, we wished to establish commonalities and differences between the distantly related apicomplexan parasites, Toxoplasma and Plasmodium, with the hope that these common host proteins co-opted by these parasites might be used by other protozoa such as the poorly studied, biodefense class B parasite Cryptosporidium parvum.

Ultimately, the concept of inhibiting host factors necessary for parasite growth or optimizing innate antiparasitic molecules may prove an effective therapeutic strategy for multiple intracellular parasites, as inhibition of these targets will necessarily limit drug resistance. We hope that one or more host proteins will emerge as novel therapeutic targets or provide the basis for novel therapeutics that will drive antiparasitic drug development.

Targeting host molecules required for parasite life cycle progression:

Can we beat them at their own game?

An assessment of the parasite life cycle allowed us to identify necessary interactions with the host, without which the parasite would be unable to proliferate: it is clear that the parasite interacts with the host upon invasion, to obtain nutrients, and ultimately to lyse and leave the host cell. As the process of parasite egress was not well characterized 120 relative to the process of invasion, we believed that this process likely provided a wealth of untapped resources for antiparasitic drug development: they only had to be characterized. This act of egress is thought to take place in seconds at the end of the

48-hour erythrocytic cycle through the complete destruction of the host erythrocyte.

From our prior work demonstrating that parasites require human calpain [15], we hypothesized that host calpain is used to destabilize the erythrocyte membrane. In this work, we endeavored to provide a more global understanding of changes that occur to the host erythrocyte membrane proteome during parasite egress and furthered identified a complex GPCR-driven host pathway necessary for cytoskeletal compromise prior to parasite release.

Together, these data suggest that parasite egress does not occur as a single explosive event just at the end of the 48-hour life cycle. More accurately, the distinct events of cytoskeletal dismantling, which we now demonstrate, suggest a systematic preparation for egress that takes place over at least 15-20 hours in which the parasite gradually disrupts the cytoskeleton to ultimately allow for facile escape. Though this work represented a concentrated effort, we submit that these results do not provide an exhaustive identification of host protein involvement in parasite growth. A combination of genetic, proteomic, biochemical, and in vivo approaches will likely be required to systemically identify and characterize host proteins required for parasite life cycle progression through invasion, replication, and egress:

1. Genome-wide screens to define host genes involved in T. gondii growth.

Aside from our data implicating multiple host proteins during parasite egress

(Chandramohanadas et al., 2009) and host actin polymerization (Gonzalez et al., 2009) during parasite invasion no host proteins have been identified to be required for 121 intracellular parasite replication or growth. To determine additional host factors involved in this process using an unbiased approach to identify host genes important for parasite growth, systematic knockdown using should be used.

2. Targeted siRNA screen for host proteins involved in parasite invasion.

Though we have made significant strides in the study of host proteins that function in parasite exit, little is known about host proteins involved in parasite invasion. In fact, it has historically been assumed that the host cell remains quiescent during this process, as much attention has been paid to parasite-derived proteins that function in invasion.

3. Quantitative proteomics studies to discover host proteins that change in response to T. gondii infection.

As a complementary approach, quantitative proteomics studies will allow the determination of host proteins that change in abundance during parasite infection. This approach will involve the use of SILAC-labeling and mass spectrometry-based sequencing (Ong et al., 2002) in host cells to discover new host-parasite interactions.

Proteins that significantly change in abundance (>3 fold change) up or down may represent host proteins modified directly or indirectly by the parasite to allow for progression of its life cycle.

4. Assessing the role of host proteins during parasite infection in vivo.

While we have demonstrated that calpains have a key role in parasite egress in cell culture models (Chandramohanadas et al., 2009), it is important to explore the in vivo physiological importance of the GPCR and calpain-mediated egress pathway using mouse models of parasite infection. Validation of this pathway in animal models would

122 provide an important proof of principle to justify future therapeutic development.

Importantly, although calpain-2 is essential for embryonic development, pharmacological inhibition of calpain-1/2 in adult mice was well tolerated (Mamoune et al., 2003) and genetic ablation of both calpain-1 and calpain-2 using tissue-specific or regulated conditional CAPN4 knockout approaches have resulted in viable mice with subtle physiological defects (Kashiwagi et al.; Shimada et al., 2008; Tan et al., 2006;

Wernimont et al.). Furthermore, knockout approaches involving multiple PKC family members have been unsuccessful. Collectively, these observations bode well for the potential clinical use of selective and effective inhibitors, as well as conditional knockout systems.

Mice that are either heterozygous or homozygous knockouts of any of host gene

“hits” from the RNAi screen should be challenged with both T. gondii, P. yoelli, and P. berghei ANKA. The mouse malaria model P. yoelii is useful as this species models severe non-cerebral malaria, the time course of infection is short, and the outcome

(survival vs. lethality) is definitive. The P. berghei ANKA mouse model, in contrast, models cerebral malaria, allowing for differential investigation of host protein involvement in different models of disease. The effect of genetic depletion of screen hits on parasite infection should be compared with that achieved using standard anti-malarial drugs including artemisinin and atovaquone, as well as that seen using specific inhibitors.

PF4 and Small molecule mimics of host defense peptides: Can we optimize an evolutionarily-tuned host defense? In addition to identifying host proteins exploited by the parasite for life cycle progression, we were interested in characterizing host proteins that naturally act as antiparasitics via exploitation of a fundamental host-parasite interaction. 123

In this work we identified the human HDP protein, PF4, as an antiparasitic molecule that kills Plasmodium parasites by lysing the parasite DV, a vital parasite organ responsible for digestion of host hemoglobin. Targeting the lysosome-like vesicles of protozoan parasites may be an evolutionarily selected mechanism for mammalian hosts to kill Plasmodium and other parasites (i.e. T. brucei). How proteins such as PF4 enter host cells remains an open question, though uptake of hPF4 into the host erythrocyte seems to diminish following parasite DV lysis and parasite death, suggesting that perhaps an active, parasite-mediated mechanism allows entry of these HDP molecules into the erythrocyte cytoplasm. Since only infected cells are targeted, and parasite viability is important for uptake, this suggests that perhaps a parasite-derived or activated endocytosis-like mechanism could allow for large molecules such as PF4 to enter, while perhaps channels or pores could be responsible for small molecule uptake

(Baumeister et al., 2006; Elford et al., 1990; Kirk and Horner, 1995).

We believe that human PF4 likely has a natural role in controlling blood stage parasite growth, and that its low hemolytic potential and selective targeting of the parasite DV endow it with properties that are well tolerated in the bloodstream. However, it is unlikely that PF4 would make a useful therapeutic for malaria treatment due to the cost of goods, and potential of inflammatory damage through its chemokine domain during cerebral malarial. Thus screening for smHDPs with a similar mechanism of action may avoid the issues of PF4 while preserving its unique mechanism of action and safety; as such, smHDPs may represent a new class of small molecules with effective therapeutic potential for combating this human disease where efforts are currently challenged with rampant drug resistance. In order to optimize these molecules as potent antiparasitics, it will be necessary to understand the unique properties of parasite 124 membranes that engender this specificity and the PF4 domains responsible for this antiparasitic activity, as well as resistance studies, development of analogues, and in vivo studies:

5. Testing PF4 against model liposomes based on the composition of parasite digestive vacuolar membranes. To determine why PF4 selectively disrupts the digestive vacuole membrane causing leakage of digestive vacuole contents via direct membrane disruption, model liposomes with similar lipid content to the parasite digestive vacuole membrane should be prepared prior to assessment of membrane compromise upon PF4 treatment. The composition of parasite digestive vacuole membranes can be determined via the isolation of this organelle from whole parasites

(Goldberg et al., 1990) and analysis of lipid content within these purified preparations via

Liquid Chromatography/Mass Spectrometry (LCMS) (Ham et al., 2004).

6. Understanding the basis of selective entry of PF4 into infected erythrocytes. Our preliminary data shows that only infected erythrocytes take up PF4 and that pretreatment with protamine sulfate blocks both entry and activity of PF4 against P. falciparum. This allows us to hypothesize that initial interaction between PF4 and the parasitized erythrocyte is electrostatic and involves binding to negatively charged molecules on the surface. Considering that the erythrocyte surface proteins such as the glycophorins are highly decorated with sialic acid, we can directly test whether sialic acid is involved in the initial binding events by specifically stripping it off using neuraminidase.

7. Generating PF4-resistant parasites. Since HDP-mediated cell death via membrane perturbation is not reliant upon a specific receptor or intracellular protein

125 target, cellular resistance to such compounds should be difficult to generate. In order to test this hypothesis, P. falciparum lines that are resistant to PF4 should be attempted to be generated via serial passaging in dilutions of our experimentally determined PF4 IC50

(0.25 x IC50, 0.5 x IC50, and the full IC50 concentration). As a control, parallel cultures should also be exposed to 0.5x IC50 concentration of pyrimethamine, a well-established antiparasitic for which resistance has been reported. This should be carried out over a time period of at least 6 months. If resistant lines are successfully generated, tiling microarrays should be used to determine any genes that contribute to resistance.

Resistance would indicate that PF4 might not be acting solely via membrane disruption.

Conversely, if parasites fail to generate resistance to these compounds, this class of compounds may have potential as antimalarials that may elicit minimal parasite resistance.

8. Assessing the in vivo role of PF4 during parasite infections in genetically modified mice.

PF4 (CXCL4) belongs to the CXC family of chemokines that has a putative HDP domain at the C-terminus. PF4 can be released upon activation from alpha-granules of blood platelets in very high physiologic concentrations, locally up to 100 mM (Lambert et al.,

2007b). Until recently, CXCL4 gene expression was thought to be restricted to megakaryocytes (Brandt et al., 2000), but other cell types may produce these chemokines as well (Schaffner et al., 2005). The Poncz laboratory was the first to develop PF4 knockout mice (Eslin et al., 2004b) and also transgenic mice that overexpress human PF4 (hPF4+; (Zhang et al., 2001)). These mice are valuable tools for the study of PF4 in vivo in several disease models including thrombus formation, atherosclerosis, megakaryopoiesis, liver fibrosis, angiogenesis and tumor formation. In

126 this proposal we aim to use these mice to understand the in vivo relevance of platelet secreted PF4 in both the severe anemic model of malaria using P. yoelii (which causes disease in the peripheral vasculature and organs) versus a model of experimental cerebral malaria caused by P. berghei ANKA (which causes pathology in the brain of mice).

9. Identification of antiparasitic domains within PF4. PF4 is comprised of three domains: an N-terminal cytokine domain, a central domain responsible for tetramerization, and a C-terminal amphipathic helical region with HDP activity (C12). We have evidence that C12 retains the ability to kill parasites, at reduced potency, and that 2 of 4 lysines on the positive face of this peptide are required for activity. Using this information we can hypothesize that these properties will also be present in the full length protein. We hypothesize that the need for 2 positive charges to maintain facial amphipathicity in the α-helical domain of PF4 will also be true in the full-length protein.

To test this, we could recombinantly express full-length PF4, mutating all lysines individually and in combinations of doubles, triples, and a quadruple mutant to determine not only if two lysines are necessary for HDP activity, but also if specific lysines are important. The resultant proteins will then be tested against P. falciparum in culture, and activity of the PF4 mutants will be compared to wild type PF4 to assess for IC50 trend and antimalarial activity.

10. Development of peptidomimetics based on the HDP domain of PF4. In an effort to further understand the interaction of PF4 antimalarial activity and generate tools for use for in vivo mouse malaria studies, we will synthesize an array of derivatives based on the C12 domain of PF4 via standard synthetic routes. The underlying principle to the generation of these C12-based tool compounds is to increase their stability and 127 potency in vitro and in vivo while maintaining low host toxicity. Ultimately, this will build the knowledge for future efforts to produce a biological therapeutic for malaria.

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