Analysis of CDPK1 targets identifies a trafficking adaptor complex that regulates microneme exocytosis in Toxoplasma By Alex Wai Chan B.S. Molecular, Cell and Developmental Biology, University of California, Los Angeles, 2017 Submitted to the Biology Graduate Program in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY IN BIOLOGY at the MASSACHUSETTS INSTITUTE OF TECHNOLOGY June 2023 © 2023 Alex Wai Chan. This work is licensed under a CC BY-SA 2.0. The author hereby grants to MIT a nonexclusive, worldwide, irrevocable, royalty-free license to exercise any and all rights under copyright, including to reproduce, preserve, distribute and publicly display copies of the thesis, or release the thesis under an open- access license. Signature of Author............................................................................................................. Alex Wai Chan Biology Graduate Program Certified by......................................................................................................................... Sebastian Lourido Associate Professor of Biology Accepted by....................................................................................................................... Mary Gehring Associate Professor of Biology Member, Whitehead Institute Director, Biology Graduate Committee 1 Analysis of CDPK1 targets identifies a trafficking adaptor complex that regulates microneme exocytosis in Toxoplasma By Alex Wai Chan Submitted to the Biology Graduate Program on May 17th, 2023, in partial fulfillment of the requirements for the degree of Doctor of Philosophy at the Massachusetts Institute of Technology. ABSTRACT Apicomplexan parasites use Ca2+-regulated exocytosis to secrete essential virulence factors from specialized organelles called micronemes. Ca2+-dependent protein kinases (CDPKs) are required for microneme exocytosis; however, the molecular events that regulate trafficking and fusion of micronemes with the plasma membrane remain unresolved. In this thesis, I describe the discovery and characterization of a regulator of microneme exocytosis in Toxoplasma gondii. In the first chapter, I introduce T. gondii as a model apicomplexan to study motile stages during asexual stages. In the second chapter, I discuss combining sub-minute resolution phosphoproteomics and bio- orthogonal labeling of kinase substrates in T. gondii to identify 163 proteins phosphorylated in a CDPK1-dependent manner. In addition to known regulators of secretion, I identify uncharacterized targets with predicted functions across signaling, gene expression, trafficking, metabolism, and ion homeostasis. In the third chapter, I describe the functional characterization of a target of CDPK1, the putative activating adaptor HOOK. In other eukaryotes, HOOK homologs form the FHF complex with FTS and FHIP to activate dynein-mediated trafficking of endosomes along microtubules. I show the FHF complex is partially conserved in T. gondii, consisting of HOOK, an FTS homolog, and two parasite-specific proteins (TGGT1_306920 and TGGT1_316650). CDPK1 kinase activity and HOOK are required for the rapid apical trafficking of micronemes as parasites initiate motility. Moreover, parasites lacking HOOK or FTS display impaired secretion of microneme proteins , leading to a block in the invasion of host cells. Taken together, our work provides a comprehensive catalog of CDPK1 targets and reveals how vesicular trafficking has been tuned to support a parasitic lifestyle. Thesis Supervisor: Sebastian Lourido Title: Associate Professor 2 ACKNOWLEDGEMENTS I would like to thank the individuals that have supported my scientific journey. Dr. Alvaro Sagasti and Dr. Sue Biggins for allowing me into their laboratories and supporting me long after my time there. I owe my motivation to pursue a career in biological discovery to my experiences in their labs. The MIT Biology program, especially Betsey, for keeping me on the logistical train tracks to graduate even when I missed important deadlines. The MIT Biology Class of 2017 for filling my first year in New England with many fond memories in and outside the classroom. The “MIT G1 Biograd Halloween Video 2017” sets the industry standard for trainee-produced Halloween videos. My friends inside and outside MIT, especially Sheena, Sam, Chris, Skylar, Tammy, James, Sebastian, Juliet, Dianne, Danbee, and Carol for being my home away from home. My thesis committee – Dr. Douglas Lauffenburger, Dr. Iain Cheeseman, Dr. Angelika Amon and Dr. Jeffrey Dvorin– for their support and complete scientific honesty that guided my doctoral research. The undergraduate trainees that I have had the immense pleasure to work with: Nicole, Elena, and Sundeep. I am in awe of your curiosity and talent and look forward to seeing what the future holds for you. The Lourido Lab, both past and present – Clare, Saima, Diego, Benedikt, Bory, Liz, Ben, Tyler, Emily, Alice, Chris, Dylan V, Haley, Aditi, Michelle, Dominic, Dylan M, Chinmay, Justin, Faye, Aarti and Eden. Each of you has contributed to my development as a scientist. Thank you for entertaining my wild ideas and always being so generous with your time and wisdom. You made all the early mornings and late nights worth it. Dr. Sebastian Lourido for training me to be a scientist that values patience, precision, accessibility, perseverance, and compassion. From incorrectly used en dashes, fickle mass spectrometers, and failed experiments, I’m grateful for all the challenges you’ve helped me through. To current and future Lourido Lab members – tough love doesn’t feel great in the moment, but more often than not, it’ll be for the best. To my family – You have all made great sacrifices to help get me to this point, on track to get a PhD from MIT. Thank you for being a constant source of love and support. 3 TABLE OF CONTENTS Abstract 2 Acknowledgments 3 Table of contents 4 Chapter 1 – Introduction Pathogenesis and life cycle of the apicomplexan Toxoplasma gondii 6 The subcellular structures required for the motile stages of T. gondii 11 Regulation of tachyzoite motile stages 18 Role of Ca2+-dependent protein kinases in T. gondii lytic cycle 25 Chapter 2 - Identifying the substrates of CDPK1 in T. gondii Introduction 34 Sub-minute resolution phosphoproteomics 38 Myristoylation modulates CDPK1 activity and alters its interacting partners 41 Bioorthogonal labeling of direct substrates of CDPK1 46 Substrates of CDPK1 48 Discussion 50 Methods 54 Author contributions 68 Chapter 3 - The HOOK complex promotes microneme exocytosis to enable invasion Introduction 68 HOOK regulates invasion 69 The HOOK complex is required for microneme exocytosis 75 HOOK regulates microneme trafficking during parasite motility stages 77 Discussion 80 Methods 83 Author contributions 92 Chapter 4 – Conclusions 94 References 98 Appendices 118 4 CHAPTER 1: Introduction Diversity within the eukaryotic tree of life is not dominated by multicellular creatures such as animals and plants, but is overwhelmingly populated by microscopic unicellular organisms referred to as protists. The SAR (stramenopiles, alveolates, and Rhizaria) supergroup houses approximately 3/5ths of eukaryotic diversity exhibiting a broad range of survival strategies that include free-living autotrophs and obligate intracellular heterotrophs (Burki 2014). These strategies are enabled by the evolution of diverse cellular morphologies. Within Alveolata, Apicomplexa are a phylum of obligate intracellular parasites that infect a wide range of animal hosts. These include several apicomplexans that cause human diseases such as malaria (Plasmodium), fatal diarrhea (Cryptosporidium), babesiosis (Babesia) and toxoplasmosis (Toxoplasma). Transmission and survival of these pathogens requires developmental adaptations across distinct host environments during their life cycle. The parasitic life cycle generally spans definitive hosts where sexual reproduction occurs and intermediate hosts where asexual reproduction occurs, although a subset displays a monoxenous life cycle. While the apicomplexan intermediate hosts of coccidia and haemosporidia are predominantly vertebrate animals, definitive hosts vary from mosquitoes (Plasmodium), ticks (Babesia), canines (Neospora) to felids (Toxoplasma). Apicomplexans have developed distinct cellular mechanisms required for their life cycles, but share conserved subcellular features that enable them to traverse intracellular and extracellular environments within their hosts. In this introduction, I describe the life cycle and pathogenesis of T. gondii, which is used as a model to study apicomplexan biology. Next, I describe the conserved subcellular features—collectively referred to as the apical complex—that enable motility- mediated pathogenesis. Then, I explain the molecular regulation of the apical complex that enables parasite motility. Finally, I will discuss the role of Ca2+-dependent protein kinases in regulating motility. 5 Lourido, S. Trends in Parasitology. 2019. Introductory Figure 1. The life cycle of Toxoplasma gondii. Sexual reproduction occurs within the intestines of felids and produces infectious oocysts. Infectious oocysts infect intermediate hosts and transition to asexual stages of the parasite that include tachyzoites for acute infection and bradyzoites for chronic infection. Acute infection involves repeated rounds of replication and lysis in virtually any nucleated cell type. Differentiation of tachyzoites to bradyzoites that form tissue cysts and chronic infection are comparably slower replicating and persist for the lifetime of the host. Consumption of tissues cysts by felids completes the life cycle of T. gondii. Pathogenesis and life cycle of the model apicomplexan Toxoplasma gondii As the definitive hosts of T. gondii, infected felids harbor the sexual reproduction of T. gondii within their intestines to produce infectious oocysts that are excreted in their feces. Oocysts are extremely environmentally resistant and were reported to be infectious in mice even after 12 months in a feces solution suspended in 33% zinc sulfate 6 and tap water (Introductory Figure 1) (Hutchison 1965). Infection of intermediate hosts occurs after consumption of food or water contaminated with oocysts. Within the intermediate host, oocytes undergo a transition from sexual to asexual stages and disseminate throughout the organism, as asexual-stage parasites can infect virtually any nucleated cell type. Asexual stages are defined by two stages that interconvert: the acute infection with tachyzoites that undergo rapid rounds of replication and lysis and chronic infection with slow-replicating bradyzoites that form tissue cysts (Introductory Figure 1). Felid predation of intermediate hosts infected with bradyzoite tissue cysts completes the life cycle as differentiation to sexual stages to produce oocyst occurs (Introductory Figure 1). Intermediate hosts include warm-blooded animals. The first documented animal fatality by toxoplasmosis was reported in a 4-month old dog that succumbed to acute infection in 1910 (Weiss and Kim 2020). Investigation into mysterious abortions in agriculturally important sheep in New Zealand led to the discovery of acute infection with tachyzoites in the late 1950s (Hartley and Marshall 1957; Hartley 1961, 1964). The first foodborne identification of T. gondii were from infected porcine in 1955 (Sanger and Cole 1955). Marine mammals and birds are also susceptible to infection (Dubey et al. 2010). The first human T. gondii infection was reported in a newborn in 1938 that suffered from symptoms of congenital acute infection including encephalitis and retinochoroiditis (Wolf, Cowen, and Paige 1939a, [b] 1939). Post-mortem analysis identified asexual-stage parasites in the eyes and central nervous system, which remained infectious when isolated parasites were used to infect rabbits and mice. Approximately 25% of the global human population has been infected with T. gondii. While seroprevalence is estimated to be approximately 11% in the United States, this can exceed 90% in some countries (Montoya and Liesenfeld 2004). Transmission of Toxoplasma to humans occurs through various routes. Infections occur through ingesting contaminated food or water that contain oocysts or food containing asexual stage tissue cysts called bradyzoites. Ingestion of undercooked meat was formally demonstrated as a route of transmission through the experimentation with 7 children in a Paris sanatorium in the 1960s, in which children were fed barely cooked beef or horse meat which led a rise in T. gondii seropositivity rates from 10% to 50% and increased to 100% after the introduction of barely cooked lamb (Desmonts et al. 1965). Fecal-oral transmission was demonstrated that same year in mice infected with oocysts derived from cat feces at the University of Glasgow (Hutchison 1965). Congenital transmission can occur in acutely infected mothers either through primary exposure to oocysts or tissue cysts or through recrudescent infection in immunocompromised women where tissue cyst bradyzoites convert to tachyzoites and resume acute infection (Mitchell et al. 1990; Montoya and Liesenfeld 2004). Recrudescence can also occur in other immunocompromised individuals such as those suffering from HIV/AIDS or those receiving immunosuppressive therapies (Mitchell et al. 1990; Montoya and Liesenfeld 2004). T. gondii infection of immunocompetent individuals is predominantly asymptomatic, but life-threatening disease can occur in the developing fetus or immunocompromised individuals. Congenital infection may result in abortion, developmental delay, and recurrent chorioretinitis (Jones et al. 2001; Torgerson and Mastroiacovo 2013). Encephalitis, pneumonia, and multi-organ failure have also been observed during T. gondii infection (Montoya and Liesenfeld 2004). Immunocompromised individuals such as those with HIV/AIDS are administered pyrimethamine and sulfadiazine Introductory Figure 2. The lytic cycle of asexual tachyzoites. Acute infection is characterized by replicative and motile stages of tachyzoites. Active invasion forms a replicative niche enabling replication. After replication is complete, motile stages are engaged to egress, initiate a unique form of cellular movement called gliding, and invade a new host cell. Cytosolic Ca2+ concentrations increase during motile stages of the tachyzoites. 8 prophylaxis to prevent recrudescent infection and prevent pneumocystis infection (Montoya and Liesenfeld 2004). Intermediate host infection involves acute infection with tachyzoites and chronic, life-long infection with bradyzoites (Introductory Figure 1). Tachyzoites are two-by- seven micron parasites that actively invade a host cell. A parasitophorous vacuole is derived from the host cell membrane that serves as a distinct replicative niche (Mordue and Sibley 1997). Replication within the vacuole results in cell doubling every 6-8 hours and lysis of the host cell occurs after 24-48 hours by engaging motile stages of the parasite (Introductory Figure 2) (I. J. Blader et al. 2015). Innate immune responses mediated by IFN-gamma results in clearance of tachyzoites within weeks of acute infection (Dupont, Christian, and Hunter 2012). Before tachyzoite infection is cleared, differentiation of tachyzoites to bradyzoites enables the formation of tissue cysts that can persist for the lifetime of the host. Tissue cysts can be found in the brain and muscle tissue and are composed of hundreds of slow-replicating bradyzoites that infrequently lyse the host cell when compared to tachyzoites (Dubey, Lindsay, and Speer 1998; Watts et al. 2015). Bradyzoites cannot be cleared by the immune system and no treatments for chronic infection currently exist. Recrudescent infection in an immunocompromised host originates from conversion of bradyzoites back to tachyzoites to resume acute infection. T. gondii has several features that facilitate experimentation. T. gondii can be cultured in primary human host cells. A majority of Toxoplasma research has been performed on a strain isolated in 1937 that was named after the initials of its child host— R.H.—who died from injuries from a baseball bat unrelated to the infection (Sabin 1941). RH parasites were maintained for years in mice and have lost their ability to form oocysts (Dubey and Hoover 1977). Currently, T. gondii parasites are typically maintained in primary human foreskin fibroblasts where several advances have contributed to its tractability as a model. First, exogenous DNA can be introduced through electroporation- mediated transfection (Soldati and Boothroyd 1993). Second, selectable markers facilitate isolation of genetically modified parasites (R. G. Donald and Roos 1993; R. G. Donald et al. 1996; Kim, Soldati, and Boothroyd 1993). Efficiency of genetic engineering 9 in haploid parasites was significantly improved by generating strains deficient in non- homologous end-joining, thus biasing repair mechanisms to homologous recombination (Fox et al. 2009). Lastly, CRISPR/Cas9 technology was adapted to tachyzoites enabling targeted methods to generate double-stranded breaks to introduce DNA of interest or to generate loss-of-function mutations (Shen, Brown, et al. 2014; Sidik et al. 2014). Of the 8,637 genes encoded in the T. gondii GT1 genome, 4,342 currently have no annotated functions and the majority of the remaining genes are annotated based on sequence homology and lack experimental validation (ToxoDB). Pooled forward genetic screens have enabled the experimental derivation of fitness scores for the 8,151 predicted protein-coding genes for T. gondii tachyzoites grown in primary human cell culture, and the approach has been adapted to more complex contexts such as screening during murine infection and the chronic stages of the parasite (Sidik, Huet, et al. 2016a; Giuliano et al. 2023; Waldman et al. 2020). While these approaches are valuable in determining the genetic requirements for T. gondii at scale, they are largely agnostic to critical information about the spatiotemporal functions of proteins including protein localization, interactions, temporal regulation, and post-translational modifications. By contrast, candidate-based approaches have enabled these in-depth analyses of proteins on a gene-by-gene basis, but require significant time and labor making the approach difficult to scale genome wide. In T. gondii, forward-genetic screening has been modified for complex phenotyping of a subset of the kinome during tachyzoite stages. This was accomplished by generating a tagging vector functionalized with a degron and fluorescent reporter enabling temporal control of protein stability and localization, respectively (Smith et al. 2022). Tagging of the T. gondii kinome followed by screening in arrayed and pooled formats resulted in 40 new kinase localizations and 15 kinases with previously uncharacterized functions during replicative and motile stages (Smith et al. 2022). Tagging-based screens begin to bridge the gap between the scale of forward-genetics and rich information obtained from phenotypic assays, but currently cannot be applied to a library of more than a couple hundred genes in T. gondii. Complementary approaches are needed to facilitate characterization of functional 10 relationships between proteins that will enable rapid integration of factors into a systems level understanding of cellular processes. The motile stages of tachyzoites which include host cell egress, gliding, and invasion require the regulated exocytosis of apicomplexan-specific organelles that is dependent on Ca2+ signaling within the parasite (Lourido and Moreno 2015). The regulated secretion of virulence factors from these organelles, called micronemes and rhoptries are required for motile stages, however, many of the molecular mechanisms mediating the trafficking and fusion of these organelles remain unidentified. In the next sections, I will describe the identity and function of the subcellular structures that are required for parasite motility and then describe the known pathways that regulate Ca2+- stimulated exocytosis. Finally, I will explain the importance of Ca2+-dependent protein kinases (CDPKs) in transducing Ca2+ signals. As the downstream substrates of these kinases are likely contributing to Ca2+-regulated exocytosis, these findings motivate the identification and characterization of CDPK1 targets. The subcellular structures required for the motile stages of T. gondii The motile stages of the parasite are required for the obligate intracellular parasitic lifestyle of T. gondii as they are crucial to the organism’s survival and spread. The motile stages are characterized by three behaviors in tachyzoites: host cell egress, invasion, and gliding (Introductory Figure 2). These processes are exquisitely coordinated by the divergent cytoskeletal and secretory components collectively referred to as the apical complex, the defining feature of the apicomplexan phylum (Introductory Figure 3). Invasion of a host cell is an active process and differs from other intracellular pathogens that induce host membrane ruffling or actin reorganization (Morisaki, Heuser, and Sibley 1995). Active penetration of the host cell plasma membrane results in the formation of a specialized compartment derived from the host called the parasitophorous vacuole (PV) (Suss-Toby, Zimmerberg, and Ward 1996). Parasites replicate within the PV–which remains a distinct compartment that does not acidify nor fuse with other membrane compartments of the host cell(Mordue et al. 1999). Once replication is complete, the 11 parasite exits the host cell by simultaneously permeabilizing the PV and initiating gliding motility, resulting in the lysis of its host cell. These processes rely on the regulated exocytosis of specialized secretory organelles called micronemes and rhoptries and the coordination of actin- and tubulin-based structures. In this section, I describe the subcellular structures that comprise the apical complex and how these components function to promote motile stages of tachyzoites. Tubulin-based cytoskeleton: the conoid, intraconoidal microtubules, and subpellicular microtubules T. gondii tachyzoites have an intricate network of tubulin- based cytoskeletal structures that are essential for both replicative and motile stages, but this Introductory Figure 3. Schematic of a T. gondii tachyzoite with subcellular discussion will focus on those relevant for structures labeled. motility. Tachyzoite dimensions are approximately 7 µm long and 2 µm wide and are highly polarized as defined by the apical complex. The conoid is located at the apical end of the parasite and is required for invasion (Carey et al. 2004). It is composed of 10-14 non-tubular filaments that are about 430 nm long and are bundled into a left-handed helical arrangement forming a barrel with a 380 nm diameter (Hu, Roos, and Murray 2002; N. S. Morrissette, Murray, and Roos 1997; Nichols and Chiappino 1987). The conoid is connected to two preconoidal rings that are connected to two 400 nm long intraconoid microtubules that extend into the cell body. In intracellular parasites, the apical polar ring (APR) is the most apical tubulin-based structure and is connected to the conoid opposite to the preconoidal rings, however, the the complex of conoid, preconoidal rings, and intraconoid 12 microtubules can extend beyond the APR when motile stages are engaged in a process called conoid extrusion. Conoid extrusion is dependent on an increase in cytosolic Ca2+ and is required for invasion (Katris et al. 2014). Recent studies have demonstrated an additional function of the preconoidal rings in regulating glideosome function, the actomyosin force generation structures that promote gliding. The actin nucleator Formin1 is recruited to the preconoidal rings to regulate F-actin formation at the conoid enabling myosin H-mediated extrusion of the conoid (Dos Santos Pacheco et al. 2022; Tosetti et al. 2019). The characteristic crescent-shaped morphology of the parasite is partially conferred by the subpellicular microtubules which originate from the APR and span approximately two-thirds of the parasite length. There are 22 cortical microtubules spaced approximately 20 nm apart that radiate out from the APR in a helical arrangement and associate below the inner membrane complex (Nichols and Chiappino 1987; Naomi S. Morrissette and David Sibley 2002). Cortical microtubules have their minus end oriented at the apex and do not display the typical growth and catastrophe dynamics observed in other systems, but are highly stable and can withstand cold and detergent treatments (Leung et al. 2017). The conoid, intraconoidal microtubules, preconoidal rings, APR, and cortical microtubules not only contribute to the structural integrity of the parasite, but also surround critical secretory organelles, which include the micronemes and rhoptries discussed below. Secretory organelles: micronemes Micronemes are specialized secretory organelles that are exocytosed to release proteins required for egress, gliding, attachment and invasion. Micronemes are small missile-shaped, approximately 50 nm in diameter, electron dense membrane bound organelles that localize to the apical end of the parasite (Introductory Figure 3) (Dubremetz and Ferguson 2009). There are approximately 50-100 micronemes in a single tachyzoite, but the number can vary depending on the life cycle stage (Vern B. Carruthers and Tomley 2008). They have been associated with the cortical microtubules in T. gondii and Plasmodium and microtubule-based trafficking of micronemes has been proposed, but more centrally located micronemes have been observed suggesting they are not all 13 microtubule associated (Leung et al. 2017; Bannister et al. 2003). Subpopulations of micronemes have even been proposed within those localized to the apex. Overexpression of the RAB5A or RAB5C GTPases resulted in the loss of laterally localized micronemes while those in the conoid remained intact (Kremer et al. 2013). While disrupting cortical microtubules resulted in microneme mislocalization throughout the cytosol, the molecular mechanisms that link the cargo to cortical microtubules remains unknown(Leung et al. 2017). Artificially raising intracellular Ca2+ by treating parasites with an ionophore is sufficient to stimulate microneme exocytosis and triggers the rapid trafficking of the organelles into the apical complex (V. B. Carruthers, Moreno, and Sibley 1999; Sidik, Huet, et al. 2016a). Based on the cortical microtubule polarity, this is consistent with minus-end directed trafficking of micronemes by the conserved motor dynein. Many members of the dynein-dynactin complex can be identified by sequence homology, but only the dynein light chain type 1 members have been experimentally characterized revealing that DLC8a localizes to the conoid and is required for microneme exocytosis and rhoptry positioning (Qureshi et al. 2013; Hu et al. 2006; Lentini et al. 2019; Gordon and Sibley 2005). Major questions remain on the experimental identification and characterization of the dynein complex. Eukaryotes typically encode a single dynein heavy chain yet are able to transport diverse cargos. Specific trafficking is mediated by activating adaptor complexes that mediate the linkage between specific cargo and the motor and activate movement (Reck-Peterson et al. 2018). Prior to the work presented in this thesis, no such activating adaptors have been identified in T. gondii yet. The function of the micronemes is mediated by the exocytosis of protein contents. 32 microneme proteins have been characterized, however, recent spatial proteomics designate 51 proteins as localizing to micronemes in tachyzoites (Brown, Sibley, and Lourido 2020; Barylyuk et al. 2020; Weiss and Kim 2020). To date, microneme proteins have been shown to have diverse functions during motile stages including exclusive functions for egress, gliding, attachment, rhoptry discharge, and active invasion. Microneme proteins were first discovered in Plasmodium merozoites and shown to 14 mediate attachment to host cells (Camus and Hadley 1985). They were subsequently demonstrated to be required for gliding when the absence of the TRAP protein caused defects in sporozoite motility and invasion (Sultan et al. 1997). The orthologue of TRAP in T. gondii, MIC2, was also required for tachyzoite gliding and invasion upon conditional knockout (Huynh and Carruthers 2006). Microneme proteins are translated into the ER where they undergo several post-translational modifications including glycosylation and proteolytic cleavage in the endomembrane system, which are important for their function and trafficking (Parussini et al. 2010; Dogga et al. 2017; J. M. Harper et al. 2006; Huynh et al. 2015; Brydges et al. 2008; El Hajj et al. 2008). Approximately 40% of microneme proteins contain at least one transmembrane domain and soluble microneme proteins hitchhike on transmembrane proteins in the endosomal system during trafficking (Reiss et al. 2001). Micronemes are exocytosed at the apical complex, exposing luminal domains and secreting soluble proteins that have domains enabling protein-lipid, protein-protein, and protein-carbohydrate interactions (Weiss and Kim 2020). Secretion of the perforin-like protein PLP1 permeabilizes the parasitophorous vacuole to promote egress from host cells (Kafsack et al. 2009). The exposed adhesin domains on an array of microneme proteins bind to surface proteins on host cells. These adhesins are then linked to the force-generating machinery embedded within the parasite pellicle, called the glideosome, which powers the anterior-posterior translocation of microneme adhesins that enable the gliding movements of the cell (Sibley 2003; Keeley and Soldati 2004; Soldati and Meissner 2004; Sheiner et al. 2010). Upon translocation, transmembrane microneme proteins are shed from the parasite surface by proteolysis mediated by the rhomboid proteases ROM4 and ROM5 (Buguliskis et al. 2010; Shen, Buguliskis, et al. 2014; Rugarabamu et al. 2015). While absence of ROM4 enhanced host attachment as expected, only mild invasion and gliding phenotypes were observed suggesting microneme shedding is not required during active invasion; however, shedding is partially required during mouse infection so it has been suggested that shedding may function during immune evasion (Rugarabamu et al. 2015; Lagal et al. 2010). 15 Actomyosin-mediated force generation: the glideosome The glideosome is a cytoskeleton-associated motor that generates the force required for the gliding movements during egress and invasion of a host cell. Upon exocytosis, transmembrane microneme adhesins bind to host cell receptors in the extracellular space and are anchored to the F-actin cytoskeleton that lies in between the parasite plasma membrane and the pellicle. The glideosome-associated connector (GAC), a conserved armadillo repeat protein, links the cytosolic domain of MIC2 to actin (Jacot et al. 2016). Tachyzoites contain short 50-100 nm unbranched actin filaments that are located between the plasma membrane and pellicle (Sahoo et al. 2006). The pellicle or inner membrane complex (IMC) is a patched network of thin alveolar lipid bilayer sacs located beneath the plasma membrane the tile the parasite periphery. Cortical microtubules are anchored on the cytosolic-facing side of the IMC whereas F-actin is anchored on the plasma membrane-facing side. F-actin attachment to the IMC is mediated by the glideosome protein complex composed of transmembrane and integral membrane proteins (GAP40, GAP45, GAP50), a myosin light chain MLC1 as well as a class XIV family motor myosin A (MyoA) that controls gliding (Foth, Goedecke, and Soldati 2006; Meissner, Schlüter, and Soldati 2002; Frénal et al. 2010; Opitz and Soldati 2002). Anterior translocation of microneme adhesins is powered by the activity of the glideosome which remains anchored in the IMC but translocates the adhesin-bound F- actin towards the basal end of the parasite (Introductory Figure 4). Secretory organelles: rhoptries Rhoptries are another secretory organelle located in the apical complex that are required for invasion, establishment of the vacuole membrane, and modulation of host cell transcription and immune evasion. Rhoptries have an elongated club-like shape that is generally divided into two regions: a tubular neck positioned within the apical complex and a posterior bulb positioned more within the cell body (Introductory Figure 3) (Alexander et al. 2005; Lebrun et al. 2005; Roger et al. 1988; Tetley et al. 1998). Tachyzoites contain 6-12 rhoptries that are grouped together at the apical end, but only 16 two access the inside of the conoid at any given time. The proteins within the neck are required for invasion while proteins in the neck are required establishing the vacuole and modulating the host cell’s transcription and immune response (Dubremetz 2007; Bradley et al. 2005; Counihan et al. 2013; Boothroyd and Dubremetz 2008; Bradley and Sibley 2007; Kemp, Yamamoto, and Soldati-Favre 2013; Saeij et al. 2006, 2007). Additional electron-lucent vacuoles have historically been observed anterior to the rhoptry and believed to be discharged rhoptries, however, recent cryo-electron tomography studies revealed that this is a distinct organelle called the apical vesicle that is anchored to the plasma membrane via a complex of proteins called the rhoptry secretion apparatus (Nichols, Chiappino, and O’Connor 1983; Porchet-Hennere and Nicolas 1983; Mageswaran et al. 2021). The apical vesicle and rhoptry secretion apparatus are required for proper docking and discharge of rhoptries. Apical vesicles were also observed to line the intraconoid microtubules, suggesting a potential mechanism for the re-loading of rhoptries for secretion (Mageswaran et al. 2021). Rhoptry proteins must cross multiple membranes to be secreted into a host cell: the rhoptry membrane, the apical vesicle, the parasite plasma membrane, and the host plasma membrane. The precise mechanism by which rhoptry discharge is regulated remains unclear but there has been a more recent indication that a subset of microneme proteins are required to trigger rhoptry exocytosis (Sidik, Huet, et al. 2016a; Sparvoli et al. 2022). Rhoptry proteins are generally divided into rhoptry neck proteins (RONs) and rhoptry bulb proteins (ROPs). Approximately 78 rhoptry proteins have been described, however, recent spatial proteomics data suggest there are 106 rhoptry proteins revealing a significant number of uncharacterized effectors (Weiss and Kim 2020; Barylyuk et al. 2020). Five identified proteins associate with the cytosolic face of the rhoptry and facilitate trafficking, positioning, and bundling of the rhoptries (Bradley et al. 2005; Cabrera et al. 2012; Beck et al. 2013; Mueller et al. 2013, 2016). Five identified proteins are integral membrane proteins on the rhoptry required for the positioning and ultrastructure of the organelle (Beck et al. 2013; Frénal, Kemp, and Soldati-Favre 2014; Hammoudi et al. 2018). Fifty three characterized proteins are luminal and have diverse 17 molecular functions including kinases, proteases, and moving junction proteins that function during acute and chronic infection of the parasite (Weiss and Kim 2020). I will focus on the moving junction in more detail as that is directly involved in active invasion of the host cell. The moving junction is a complex of proteins that directly connects the parasite and host cell during invasion. The parasite embeds a protein complex in the host plasma membrane that functions as a parasite-derived receptor used as an anchoring substrate to propel itself into the host cell using glideosome-generated force. RON2 is inserted into the host plasma membrane and interacts with the host cytoskeleton indirectly by recruiting RON4, RON4L1, RON5, and RON8 in the host cytosol (Lebrun et al. 2005; Besteiro et al. 2009; Straub et al. 2011). The microneme protein AMA1 is secreted onto the surface of parasites and connected to the underlying glideosome and binds to RON2 during active invasion into the host cell (Alexander et al. 2005; Lebrun et al. 2005). Rhoptry proteins not only have important signaling functions within the host, but are required to form receptors on the host plasma membrane required for active invasion. Regulation of the tachyzoite motile stages The apical complex and actomyosin system coordinate the processes that promote motility, but these events require rapid spatiotemporal control to function successfully. Tachyzoites spend most of their life cycle replicating inside a host cell— typically 24 to 48 hours—but must eventually egress and invade new host cells to survive (Introductory Figure 2). The transition from replicative to motile stages requires the initial exocytosis of micronemes for membrane rupture and gliding followed by rhoptries for invasion. This section summarizes the cellular signaling pathways that regulate this critical transition to motility. These include the coordination of cyclic nucleotide-, lipid-, and Ca2+-signaling effectors that regulate motility. When to leave: initiating the signaling cascade Extrinsic and intrinsic signals have been identified that initiate the signaling required to trigger microneme exocytosis. Three extrinsic signals have been identified to 18 trigger tachyzoite egress, but the molecular mechanisms mediating these signals remains unresolved. Disruption of the host cell membrane is sufficient to trigger parasite egress, which is dependent on a decrease in host cytosolic K+ concentrations, which is normally high (Moudy, Manning, and Beckers 2001). Low pH in the host cytosol also triggers parasite egress, which may accumulate as the parasite burden within the host cell increases (Roiko, Svezhova, and Carruthers 2014). The sensors for K+ and H+ are likely independent as low pH can overcome the inhibitory effects of high K+ that normally represses egress (Roiko, Svezhova, and Carruthers 2014). Serum albumin was also shown to stimulate egress (Brown, Lourido, and Sibley 2016). Lastly, abscisic acid was also found to stimulate egress via production of cADPR (Nagamune et al. 2008; Chini et al. 2005). While microneme exocytosis is affected by ion concentrations, serum albumin and abscisic acid, the sensors remain unidentified. Recent work has identified an intrinsic signal stimulating parasite egress, revealing phosphatidic acid as a key signaling molecule (Bisio et al. 2019). Intracellular parasites secrete the diacylglycerol kinase 2 (DGK2) outside of the parasite into the parasitophorous vacuole where it is required for the accumulation of the lipid, phosphatidic acid (PA), on the outer leaflet of the parasite plasma membrane (Bisio et al. 2019). It has been proposed that accumulation of PA on the outer leaflet is sensed by an atypical guanylate cyclase (GC) which is a multipass transmembrane protein in the parasite plasma membrane that has a P4-ATPase which are known to bind and transport phospholipids from the outer to inner leaflet (Bisio et al. 2019). Extrinsic and intrinsic signals were placed upstream in the signaling pathway as their effects are still dependent on cyclic nucleotide signaling which is upstream of intracellular Ca2+ release. Transmitting the signal: cGMP-mediated signaling While the purine cyclic nucleotide cGMP was known to be a critical second messenger in stimulating Ca2+ signaling, the guanylate cyclase responsible for cGMP production was only recently identified (Bisio et al. 2019; Yang et al. 2019; Brown and Sibley 2018). The atypical guanylate cyclase is a 477 kDa, 22-transmembrane protein that localizes to the apical cap of parasites and converts GTP to cGMP (Brown and 19 Adapted from Lourido, S. & Moreno, SNJ, Cell Calcium. 2015. Sibley 2018; Jia et al. 2017). It has an unusual P4-ATPase flippase at its N termini that is required for proper guanylate cyclase activity (Brown and Sibley 2018; Bisio et al. 2019). Loss of the cyclase blocks egress, microneme protein secretion, and low cytosolic cGMP levels—defects that can be rescued by adding in an exogenous cell-permeable cGMP analogue PET-cGMP (Brown and Sibley 2018; Bisio Introductory Figure 4. Molecular regulation Ca2+- et al. 2019; Yang et al. 2019). mediated exocytosis. cGMP-mediated activation of cGMP levels are PKG leads to an accumulation of PIP2 that is converted to second messenger IP3 and DAG. DAG is regulated by converted to phosphatidic acid which participates in phosphodiesterases (PDEs) microneme exocytosis. IP mediates Ca2+3 release from the ER, which subsequently activate CDPKs to that convert cGMP to GMP, promote microneme exocytosis. thus having an inhibitory effect on initiating cGMP-mediated signaling. Most knowledge on PDEs originates in Plasmodium, which encodes four PDEs (PDEα, -β, -γ, and -δ) that degrade cGMP (Perrin et al. 2020; Baker et al. 2017). PDEβ—which is essential in asexual blood stages—also degrades cAMP, a second messenger that antagonizes motility (Perrin et al. 2020; Baker et al. 2017). Toxoplasma encodes 18 PDEs, 11 of which are expressed in tachyzoites (Moss et al. 2022). Functional characterization of expressed PDEs through conditional knockdown approaches revealed that PDE1, 2, 5, and 9 participate in lytic cycle growth of tachyzoites, with PDE1 and PDE2 having the most severe defects including extracellular gliding for PDE1 and motile and replicative defects for PDE2 (Moss et al. 2022). The ability to control PDE activity has become an important experimental tool to 20 study tachyzoite motile stages. Treatment of intracellular tachyzoites with the cGMP- PDE inhibitors, zaprinast and BIPPO, rapidly raises intracellular cGMP by inhibiting its degradation, thus activating PKG and stimulating tachyzoite egress and microneme exocytosis (Stewart et al. 2017; Brown, Lourido, and Sibley 2016; Sidik, Hortua Triana, et al. 2016). To summarize the pathway thus far, extrinsic and intrinsic signals activate an atypical guanylate cyclase to raise intracellular cGMP levels. In the absence of these signals, cGMP levels are kept low by phosphodiesterases that can be inhibited by small- molecule inhibitors to artificially raise intracellular cGMP concentrations and activate motility. cGMP activation of motile stages is exclusively mediated by the protein kinase G (PKG) (Introductory Figure 4). PKG is a serine/threonine kinase belonging to the AGC group. Apicomplexans have a single PKG gene with three cGMP-binding domains, but coccidians (Toxoplasma, Hammondia, Neospora, and Eimeria) express two isoforms that result from alternative translation (Gurnett et al. 2002; Baker and Deng 2005). PKG-I is acylated and localized to the plasma membrane, but PKG-II lacks the acylation and is cytosolic (R. G. K. Donald and Liberator 2002; Gurnett et al. 2002; R. G. K. Donald et al. 2002). This is in contrast to Plasmodium, which exclusively expresses the cytosolic proteoform. Chemical-genetic studies engineered PKG sensitivity to ATP analogue inhibitors by making a single amino acid substitution within the ATP binding pocket. This approach, among others, were used to demonstrate the kinase’s essentiality during the lytic cycle of apicomplexans (Brown, Lourido, and Sibley 2016; R. G. K. Donald et al. 2002, 2006; R. G. K. Donald and Liberator 2002; Gurnett et al. 2002; Baker and Deng 2005; Diaz et al. 2006; Doerig 2004; H. M. Taylor et al. 2010). The small-molecule inhibitors trisubstituted pyrrole inhibitor (Compound 1) and imidazopyridine inhibitor (Compound 2) were used to selectively inhibit sensitized PKG alleles (Gurnett et al. 2002; R. G. K. Donald et al. 2006). Conditional knockdown approaches were also adapted to study PKG function. The auxin-inducible degron (AID) system enables rapid protein knockdown within minutes to hours (Nishimura et al. 2009; Brown, Long, and Sibley 2017). To generate a conditional knockdown strain, a protein of interest is tagged with a mini auxin-inducible degron (mAID; approximately 7 kDa) in parasites expressing the 21 transport inhibitor response 1 (TIR1) receptor for the plant hormone auxin 3-indoleacetic acid. Auxin treatment promotes TIR1 binding and recruitment of a SCF E3 ubiquitin ligase to the mAID-tagged target, resulting in ubiquitin-dependent proteasomal degradation (Nishimura et al. 2009). This approach was used to conditionally deplete both PKG proteoforms enabling subsequent complementation with a single isoform of PKG, demonstrating that the plasma-membrane association is essential whereas the cytosolic isoform is dispensable (Brown, Long, and Sibley 2017). This approach is valuable as it allows genetically encoded protein-level knockdown that is rapid and facilitates the functional characterization of essential genes that are refractory to loss of function mutations without requiring a small-molecule inhibitor. PKG activates downstream lipid signaling to stimulate intracellular Ca2+ release (Introductory Figure 4). PKG inhibition in Plasmodium resulted in reduced levels of phosphatidylinositol 4,5-bisphosphate (PIP2), but increased levels of PIP2 precursors (Brochet et al. 2014). The enzyme phosphoinositide phospholipase C (PI-PLC) localizes to the plasma membrane in T. gondii and its knockdown resulted parasite death as a result of aberrant cell division (Bullen et al. 2016). PI-PLC is hypothesized to catalyze the production of the second messengers diacylglycerol (DAG) and inositol 1,4,5- trisphosphate (IP ) to promote intracellular Ca2+3 release and microneme exocytosis. Chemical inhibition of diacylglycerol kinases, which convert DAG to phosphatidic acid (PA), results in defects in microneme exocytosis, egress, and invasion (Bullen et al. 2016). A PA-binding protein on the surface of micronemes called APH was also shown to be required, consistent with a link between PA production on the plasma membrane and microneme exocytosis (Bullen et al. 2016; Darvill et al. 2018). In other eukaryotes, IP3 binds to an IP3 receptor on the ER to activate Ca2+ release into the cytosol (C. W. Taylor and Tovey 2010; Seo et al. 2015). The receptor remains unidentified in T. gondii, but biochemical and pharmacological evidence supports its existence. Photoactivatable IP3 activates Ca2+ release in Plasmodium (Alves et al. 2011). Isolated microsomes from T. gondii release Ca2+ after addition of IP3, but this release is inhibited in the presence of receptor inhibitors (Chini et al. 2005). 22 Ca2+ regulation of motile stages Cytosolic Ca2+ concentrations are tightly regulated within tachyzoites. During replicative stages, cytosolic Ca2+ concentrations are maintained low (50-100 nM) as measured by ratiometric chemical fluorescent probe Fura-2 (Moreno and Zhong 1996). Spikes in intracellular Ca2+ are measured with lower-affinity probes such as Fura-5F or Fluo-4 during live microscopy of motile parasites (Moreno and Zhong 1996; Lovett and Sibley 2003). Genetically encoded calcium indicators have also been adapted in apicomplexans that enable spatial localization, noninvasive live cell imaging, and lack the loading required for fluorescent probes (McCombs and Palmer 2008; Tian et al. 2009; Zhao et al. 2011; Akerboom et al. 2013; Bassett and Monteith 2017; Sidik, Hortua Triana, et al. 2016). These tools have revealed oscillations in cytosolic Ca2+ during parasite gliding that indicated cycles of reuptake are required (Borges-Pereira et al. 2015; Stewart et al. 2017). Direct manipulation of cytosolic Ca2+ is achieved by raising it using the ionophore A23187, which equilibrates Ca2+ across membranes to induce high cytosolic concentrations (~1 µM). In contrast, cytosolic Ca2+ is lowered by treating parasites with the chelator BAPTA-AM (Tsien 1980; Vieira and Moreno 2000). Indirect approaches mentioned previously include stimulation with zaprinast or PET-cGMP that promote Ca2+ mobilization through PKG signaling (Brown and Sibley 2018). Cytosolic Ca2+ concentrations influence external and internal stores. Mobilizing intracellular Ca2+ stores is required for motile stages of the parasite, but extracellular Ca2+ has been shown to enhance invasion (Pace et al. 2014). Influx of extracellular Ca2+ is likely regulated by voltage-gated Ca2+ channels that become activated after mobilization of internal Ca2+ stores. Two voltage-gated calcium channels have been identified but remain uncharacterized (Nagamune, Beatty, and Sibley 2007). The endoplasmic reticulum contains the largest intracellular store of Ca2+, although stores can also be found in the mitochondria, acidocalcisomes, and plant-like vacuole. Within the ER, the coordination of pumps, leak channels, and ligand-gated ion channels regulates intracellular Ca2+. Inhibition of SERCA–an ATPase pump that transports Ca2+ into the ER– with the small-molecule thapsigargin results in elevated cytosolic Ca2+ that occur through putative leak channels (Moreno and Zhong 1996; Nagamune, Beatty, and Sibley 2007). 23 Several transmembrane proteins have been suggested to mediate Ca2+ leakage based predicted orthologues with mammalian systems including Bcl-2, pannexin 1, presenilins, and TRPC1 (Prole and Taylor 2011; Hortua Triana et al. 2018). Lastly, Ca2+ release from the ER to the cytosol can be stimulated with IP3 as mentioned previously, but the identity of the IP3 receptor remains unknown (Alves et al. 2011; Chini et al. 2005). Increased cytosolic Ca2+ is necessary and sufficient to stimulate microneme exocytosis, egress, and gliding of tachyzoites (V. B. Carruthers and Sibley 1999). This was demonstrated using the ionophore A23187, but only recently was it shown that cytosolic Ca2+ increases directly precede egress (V. B. Carruthers and Sibley 1999; Borges-Pereira et al. 2015). Regulatory feedback on upstream signaling factors has been proposed since ionophore treatment is downstream of DAG production which is required for microneme exocytosis. This has also been suggested by recent phosphoproteomic studies on a Ca2+-regulated kinase, CDPK3 which we discuss in the next section (Nofal et al. 2022). Interestingly, ionophore treatment does not rescue loss of PKG, indicating functions for PKG both upstream and downstream of Ca2+ release (Brown, Lourido, and Sibley 2016; Brown, Long, and Sibley 2017; Brown and Sibley 2018). This is consistent with phosphoproteomic analysis of PKG in Plasmodium that identifies downstream factors involved in actomyosin activity (Alam et al. 2015). The major Ca2+-regulated effectors controlling motility are calcium dependent protein kinases which are explained extensively in the next section. Additional Ca2+- regulated effectors have been characterized, including the C2-domain containing DOC2.1, which mediates membrane fusion in other eukaryotes and is required for microneme exocytosis in Toxoplasma and Plasmodium (Friedrich, Yeheskel, and Ashery 2010; Farrell et al. 2012). Three calmodulin-like proteins localize the apical complex and interact with MyoH to promote invasion, egress, and gliding (Long, Brown, et al. 2017). Centrin2 (CEN2) contains Ca2+-binding EF hands and are required for replication, microneme secretion and invasion (Leung et al. 2019; Lentini et al. 2019). The serine/threonine protein phosphatase calcineurin (CnA) and corresponding regulatory subunit (CnB) relocalize from the cytosol to apical complex upon Ca2+ release and are required for host cell attachment (Paul et al. 2015; Rusnak and Mertz 2000). Lastly, as 24 microneme proteins are required for rhoptry discharge, identifying a specific role of Ca2+ in rhoptry exocytosis has remained elusive. Ferlin proteins are known to bind Ca2+ and a recently discovered FER2 was shown to be required for rhoptry exocytosis but dispensable for microneme exocytosis, suggesting additional roles for Ca2+ during motile stages (Coleman et al. 2018). I have described a subset of Ca2+-regulated effectors that participate in the motile stages, but there are likely several uncharacterized effectors that remain identified. To address this challenge, the Ca2+-responsive proteome was recently identified using cellular thermal shift assays coupled with quantitative mass spectrometry identifying over two-hundred effectors (Herneisen et al. 2022). Future work will not only require characterizing additional Ca2+-regulated effectors, but also require integrating them within the motility signaling program. Role of Ca2+-dependent protein kinases in T. gondii lytic cycle The protein domain architecture and activation of apicomplexan CDPKs Ca2+-dependent protein kinases (CDPKs) are enzymes with Ca2+-stimulated serine/threonine kinase activity. The canonical architecture for CDPKs includes a N- terminal kinase domain that is linked to four EF hand domains by a short inhibitory domain, which together form the calcium activation domain (CAD)(Harmon, Gribskov, and Harper 2000; Wernimont et al. 2010, 2011). This is unlike calmodulin (CaM)- regulated kinases (CaMKs) found in opisthokonts, in which the kinase effector and the CaM regulator are separate. However, CDPKs are believed to have arisen from the fusion of a kinase domain with CaM (Billker, Lourido, and Sibley 2009; Hui, El Bakkouri, and Sibley 2015). EF hands are structurally defined by a helix-loop-helix composed of a 12- residue loop flanked by 12-residue alpha helices that together coordinate the binding of a Ca2+ ion to expose hydrophobic surfaces enabling protein-binding interactions (Lewit- Bentley and Réty 2000; Wernimont et al. 2010). In apicomplexans, EF hands are generally found in three protein families: calmodulin (CaM), calcineurin B-like (CBL), and calcium-dependent protein kinase (CDPK). 74 genes are currently annotated to encode EF hand–containing proteins in the T. gondii genome (ToxoDB). The CDPKs are the only 25 family in which EF hands are linked to effector domains, whereas Ca2+ indirectly regulates other effectors through CaM and CBL proteins. In apicomplexans, CDPKs are the primary Ca2+-responsive kinases. Activation of CDPKs can be tuned to respond to a wide range of intracellular concentrations of Ca2+ from 70-100 nM to low µM (Ingram et al. 2015; J. F. Harper, Breton, and Harmon 2004; Moreno and Zhong 1996). Full-length structures of the canonical T. gondii CDPKs CDPK1 and CDPK3 have revealed the mechanism by which Ca2+ activates the kinase domain (Wernimont et al. 2010; Ojo et al. 2010; Ingram et al. 2015). Activation by Ca2+ requires relieving autoinhibition from the CAD and for a subset of CDPKs, allosteric activation by the N-terminal lobe of the kinase domain. In the inhibited state, the catalytic site of the kinase domain is occluded by an extended alpha helix derived from the junction and a portion of an EF hand in the CAD. For CDPK1, Glu135, Asp138, and Lys338 block ATP and substrate binding by simulating a pseudosubstrate (Wernimont et al. 2010). Ca2+ binding to the EF hands is sufficient to relieve autoinhibition by reorienting the CAD 130 degrees from the catalytic site (Wernimont et al. 2010). This mechanism of autoinhibition is similar to that of CaMKs in which relief is sufficient to activate the kinase domain, but this is insufficient for activation of CDPK1. CDPK1 maintains a “ɑC-out” conformation typical of inactive protein kinases and requires the additional binding of the N-terminal lobe of the kinase to the CAD for activation (Ingram et al. 2015). Loss of a single phenylalanine in the N-terminal region is sufficient to abrogate kinase activity by preventing allosteric activation (Ingram et al. 2015). While Ca2+ is no longer required after the initial activation of CaMKs, CDPKs require the continued presence of Ca2+ for kinase activity indicating that activity is reversible upon chelating Ca2+. Diversity of CDPKs within apicomplexans CDPKs have been identified in plants, ciliates, and apicomplexans, but are absent in fungi and animals. Within apicomplexans, five groups of CDPKs are conserved amongst T. gondii, P. falciparum, and C. parvum based on phylogenetic analysis (Billker, 26 Lourido, and Sibley 2009). While the T. gondii genome encodes 14 CDPKs, only 7 are conserved with Plasmodium and Cryptosporidium (Billker, Lourido, and Sibley 2009; Hui, El Bakkouri, and Sibley 2015). The first group displays the canonical domain architecture of CDPKs that have a relatively short N-terminal extension and includes TgCDPK1 (CpCDPK1, PfCDPK4), TgCDPK3 (CpCDPK3, PfCDPK1). Several CDPKs contain motifs for N-terminal lipid modifications including N-myristoylation and palmitoylation. TgCDPK3 (PfCDPK1) and TgCDPK1 (PfCDPK4) contain these modifications which have been shown to be important for localization and function (Möskes et al. 2004; Broncel et al. 2020, 2019). The numbering between apicomplexan orthologues differs since they were historically discovered and named at different times. The second group also has the canonical domain architecture, but has longer N termini compared to the first group. This includes TgCDPK2 (CpCDPK2), PfCDPK3, TgCDPK2A and 2B (PfCDPK2, CpCDPK2a), and TgCDPK5 (CpCDPK5, PfCDPK5). Only PfCDPK2 is predicted to be acylated amongst CDPKs in this group. The other groups diverge from the canonical architecture. The third group contains CDPKs with only 3 C-terminal EF hands and include TgCDPK4A, TgCDPK4, TgCDPK8, and CpCDPK4. The fourth group has one or more EF hands located N-terminal to the kinase domain followed by three to four EF hands. These include TgCDPK9, TgCDPK6 (CpCDPK6, PfCDPK6). The fifth group contains two or more N-terminal EF hands followed by a pleckstrin homology domain and a C-terminal kinase domain. These include TgCDPK7 (PfCDPK7). The function of CDPKs during motile stages of T. gondii The motile stages of tachyzoites are defined by host cell egress, gliding motility, and invasion involving CDPK1, CDPK3 and CDPK2a (Introductory Figure 4). CDPK1 was the first CDPK to be functionally characterized in T. gondii and was demonstrated to be required during motile stages of the parasite (Lourido et al. 2010). Prior to the discovery of CDPK1’s function, kinase activity was shown to be required for parasite motile stages as treatment with the pan-kinase inhibitor KT2936 blocked host cell attachment (Kieschnick et al. 2001). Microneme protein secretion—which is required for motile stages—was also blocked with serine/threonine kinase inhibitors and this 27 inhibition was not overcome by artificially raising intracellular Ca2+ levels (V. B. Carruthers, Moreno, and Sibley 1999). Furthermore, loss-of-function mutations in CDPKs in Plasmodium resulted in defects at various developmental stages but its function in mediating motile stages was unknown. These observations motivated the pursuit of CDPKs as the transducers of Ca2+ signaling during motile stages of the parasite. CDPK1 was localized to the cytosol and its function was examined by generating a conditional knockdown allele and was demonstrated to be required for microneme exocytosis, impacting all aspects of the motile stages (Lourido et al. 2010). Furthermore, loss of CDPK1 function had no consequences on parasite replication or the biogenesis of micronemes, consistent with its role in transducing intracellular Ca2+ to initiate motility. The function of CDPK1 relies on its kinase activity as a D174A mutation produced a kinase-dead allele that phenocopied the microneme protein secretion defect observed in the conditional knockdown (Lourido et al. 2010). Identification of the myristoylated proteome in T. gondii confirmed the predicted modification at Gly2 of CDPK1 (Broncel et al. 2020). Furthermore, while loss of myristoylation led to no observable differences in CDPK1 localization, parasites suffered significant defects during the lytic cycle which is partially attributed to egress (Broncel et al. 2019). CDPK3 and CDPK2a function in a subset of motility-related Ca2+ signaling compared to CDPK1. CDPK3 localizes to the cytosolic leaflet of the plasma membrane and its localization is mediated by myristoylation at Gly2 and palmitoylation at Cys3 (McCoy et al. 2012; Garrison et al. 2012; Lourido, Tang, and Sibley 2012). Loss of either acylation modifications is sufficient to mislocalize CDPK3 (McCoy et al. 2012; Garrison et al. 2012; Lourido, Tang, and Sibley 2012). While parasites are still viable in cell culture in the absence of CDPK3, the kinase is required for microneme exocytosis specifically during ionophore-stimulated conditions (Garrison et al. 2012; McCoy et al. 2012; Lourido, Tang, and Sibley 2012). The context-specific requirements of CDPK3 has been attributed to the need for the kinase in intracellular parasites that experience higher K+ concentrations that inhibit secretion, but compensation via other pathway members in extracellular parasites (Moudy, Manning, and Beckers 2001; McCoy et al. 2012). In 28 extracellular parasites, CDPK1 inhibition blocked microneme exocytosis regardless of stimulation with ionophore or zaprinast, a PDE inhibitor that stimulates PKG activity (Lourido, Tang, and Sibley 2012). In contrast, CDPK3 was only required during ionophore stimulation but not zaprinast treatment, which led to the conclusion that PKG activity could overcome CDPK3 inhibition and suggested they share a subset of substrates (Lourido, Tang, and Sibley 2012). Furthermore, while the loss of CDPK3 is dispensable in parasites grown in cell culture, CDPK3 is important during mouse infection as mutant parasites display decreased virulence (Treeck et al. 2014). CDPK3 requirement during egress but not invasion was also observed in the P. falciparum orthologue PfCDPK5 during erythrocyte infection (Dvorin et al. 2010). Epistasis between PKG and CDPK3 has also been documented in orthologues in Plasmodium berghei (Fang et al. 2018). Further characterization of CDPK3 identified additional members of its pathway that are substrates of the kinase, including SCE1 that was shown to be a suppressor of egress, the non-canonical myosin A required for the gliding machinery, as well as additional targets that suggest regulatory feedback on intracellular Ca2+ release (McCoy et al. 2017; Garrison et al. 2012; Stewart et al. 2017; Tang et al. 2014; Gaji et al. 2015; Powell et al. 2018; Nofal et al. 2022). CDPK2a also functions during motile stages to support motility by mediating microneme discharge (Shortt et al. 2022). Conditional depletion of CDPK2a results in severe defects during the lytic cycle. Similar to CDPK3, CDPK2a was required for ionophore-induced egress but zaprinast stimulation was sufficient to overcome knockdown (Shortt et al. 2022). Gliding motility and microneme exocytosis were also impaired in the absence of CDPK2a (Shortt et al. 2022). CDPK2a functions in concert with CDPK1 as opposed to the PKG/CDPK3 axis of signaling. Parasites were sensitized to a sublethal dose of a small molecule inhibitor of CDPK1 in the absence of CDPK2a, but not CDPK3 (Shortt et al. 2022). These studies highlight the extensive interconnectedness between CDPKs and other signaling effectors in mediating parasite motility. The function of CDPKs during the non-motile stages of T. gondii 29 Additional canonical and non-canonical CDPKs have been characterized during stages largely unrelated to the motile phase of the asexual life cycle. Conditional knockdown of the non-canonical CDPK7, which has N-terminal EF hands and a pleckstrin-homology domain, resulted in cell division defects without affecting egress, motility and invasion (Morlon-Guyot et al. 2014). The cell division phenotype arose from defects in centrosome and kinetochore assembly (Morlon-Guyot et al. 2014). During the chronic stages, CDPK2 was shown to be required for bradyzoite viability but dispensable for tachyzoite growth (Uboldi et al. 2015). CDPK2 has a mostly canonical domain architecture but contains a CBM20 carbohydrate module important for recruitment to amylopectin granules required for bradyzoite carbohydrate storage. Loss of CDPK2 increased production and storage of amylopectin granules in tachyzoites that was exacerbated in bradyzoites which was proposed to negatively affect viability (Uboldi et al. 2015). Granule accumulation was also observed by blocking Ca2+ signaling using the cell permeant chelator BAPTA-AM (Uboldi et al. 2015). Non-canonical CDPKs also include pseudokinases that lack kinase activity but may mediate protein-protein interactions (Boudeau et al. 2006). Seven of these were examined during acute and chronic infection in cell culture and in mice including CDPK4, CDPK4a, CDPK4b, CDPK6, CDPK7a, CDPK8, and CDPK9 (Long, Wang, and Sibley 2016). Loss of function mutations in both Type 1 and Type 2 parasites revealed that most of these were dispensable during acute and chronic infection, even under conditions where genetic interactions between these CDPKs were probed (Long, Wang, and Sibley 2016). Loss of CDPK6 function did, however, lead to a mild plaque defect in cell culture and reduced tissue cyst burden during mouse infection (Long, Wang, and Sibley 2016). These non- canonical CDPKs were largely dispensable during acute and chronic infection, but they may have uncharacterized roles during other stages of the T. gondii life cycle. Chemical-genetic and proteomic tools to study CDPK function Deconvoluting the specific contribution of an individual protein kinase in the context of a complex signaling program is a major challenge in understanding the molecular functions of these enzymes. This section describes how chemical genetics 30 and proteomics have led to significant advances in the functional characterization of CDPKs. Chemical genetics involves the use of small-molecule ligands on genetically modified proteoforms that facilitate specific analysis of an enzyme of interest. The ATP- binding pockets of eukaryotic protein kinases can be genetically engineered to be susceptible to small-molecule inhibitors or ATP analogues enabling selective inhibition or bioorthogonal chemistries (Bishop et al. 2000). A majority of eukaryotic kinases encode a hydrophobic residue at the gatekeeper which is located within the regulatory spine of kinase domains (Hui, El Bakkouri, and Sibley 2015). Gatekeeper residues control access to the ATP-binding pocket of kinases, with larger residues conferring resistance to bulky-ATP analogues. 109 kinases are predicted to be active in T. gondii and alignments within kinase subdomain V reveals that more than 80% of protein kinases have large hydrophobic gatekeeper residues (Peixoto et al. 2010). Remarkably, CDPK1 is unique among active kinases for instead harboring a glycine as its gatekeeper (Wernimont et al. 2010; Lourido et al. 2010). As a majority of kinases have large gatekeeper residues, previous studies have typically made gatekeeper substitutions that sensitize the kinase of interest to pyrazolopyrimidine (PP) derivatives in a background where other kinases are resistant (Lopez, Kliegman, and Shokat 2014). Since CDPK1 naturally harbors an expanded ATP-binding pocket in a background of insensitive gatekeepers, selective inhibition of CDPK1 with the PP derivative 3-MB-PP1 resulted in a block in microneme protein secretion, egress and invasion (Lourido et al. 2010). Specific inhibition by 5 µM 3-MB-PP1 was confirmed by generating a resistant CDPK1 allele harboring a G128M mutation at its gatekeeper and results were also confirmed in cell-free kinase assays showing an IC50 of 0.8 nM (Lourido et al. 2010). This approach was also applied to characterize CDPK3 function in tachyzoites, but the Met gatekeeper was substituted with a Gly to confer susceptibility in a CDPK1-resistant background (Lourido, Tang, and Sibley 2012). The unique susceptibility of CDPK1 has made it an attractive drug target for T. gondii infection in which potent and selective PP analogues have demonstrated efficacy during mouse infection (Hui, El Bakkouri, and Sibley 2015; 31 Rutaganira et al. 2017). The approach has also been adapted to enable identification of kinase substrates which we discuss below. Apicomplexan protein kinases have garnered a lot of attention as they are critical signaling nodes controlling pathogenesis, yet the identification and functional characterization of their kinase targets and the integration of them within signaling networks has been challenging and ongoing effort. In the past decade, the characterization of phosphorylation within apicomplexans has evolved from descriptive to more functional efforts. Improvements in mass spectrometry and methods to enrich phosphorylated peptides enabled initial identification of the phosphorylated proteome within T. gondii tachyzoites and P. falciparum schizonts, trophozoites, and ring stage parasites (Treeck et al. 2011; Solyakov et al. 2011; Lasonder et al. 2012; Pease et al. 2013). Additional utilization of genetic, chemical genetic, and pharmacological tools enabled the ability to deconvolute kinase-dependent phosphorylation from complex signaling programs in T. gondii, P. falciparum, and P. berghei (Treeck et al. 2014; Nofal et al. 2022; Fang et al. 2017; Invergo et al. 2017; Pease et al. 2018; Blomqvist et al. 2020). These include the characterization of TgCDPK3 phosphorylation of Myosin A to promote gliding, PbCDPK4 and PbSRPK1 during male gametogenesis, and PfCDPK1, PfPK7, and PfCDPK5 during erythrocytic stages (Tang et al. 2014; Gaji et al. 2015; Invergo et al. 2017; Fang et al. 2017; Pease et al. 2018; Blomqvist et al. 2020). Chemical-genetic approaches generated analog-sensitive kinases to characterize PfPKG-dependent phosphorylation (Alam et al. 2015). These approaches were adapted for labeling and enrichment of direct kinase substrates in cell lysates for TgCDPK1 and PbCDPK4 (Lourido, Jeschke, et al. 2013; Fang et al. 2017). For CDPK1, the bulky ATP analogue N6-furfuryladenosine (kinetin)-5′-O-[3- thiotriphosphate] (KTPγS) enables selective binding to CDPK1G but not CDPKM in parasite lysates (Lourido, Jeschke, et al. 2013). Addition of Ca2+ stimulates CDPK1 kinase activity in which thiophosphate from KTPγS is transferred to kinase substrates. Thiophosphorylated proteins were enriched using two methods in parallel. In the first type of enrichment, labeled lysates were treated with p-nitrobenzyl mesylate to chemically modify thiophosphorylated proteins to enable antibody-based enrichment of 32 proteins followed by trypsin digestion for proteomic analysis. The second enrichment treated labeled lysates with trypsin first and then enriched thiophosphorylated peptides with an iodoacetyl resin that reacts with thiol groups. Captured peptides were eluted with Oxone monopersulfate as phosphorylated peptides. LC-MS/MS was performed to identify enriched peptides from both methods to generate a high confidence list of CDPK1 substrates (Lourido, Jeschke, et al. 2013). Similar experiments were performed for PbCDPK4 in Plasmodium (Fang et al. 2017). For bio-orthogonal labeling of direct substrates, previous thiophosphorylation experiments for TgCDPK1 and PbCDPK4 in parasite lysates identified fewer than 20 phosphosites (Lourido, Jeschke, et al. 2013; Fang et al. 2017). These approaches did not utilize methods to enable relative quantitation, making it difficult to directly compare results from sensitive and insensitive alleles. The dynamin related protein B (DrpB) was validated as a substrate of CDPK1 and is required for microneme biogenesis in cell culture (Lourido, Jeschke, et al. 2013; Breinich et al. 2009). Since microneme biogenesis occurs upstream of motile stages, it is unclear if this process is direct, especially since loss of CDPK1 does not affect microneme biogenesis. Thus, a major question remains on the identity of additional CDPK1 targets and their role in regulating motile stages. This thesis focuses on understanding the functions of CDPK1 by identifying the kinase’s substrates and uncovers new molecular pathways that participate in parasite motile stages. 33 CHAPTER 2: Identifying the substrates the Ca2+-dependent protein kinase 1 in T. gondii Alex W Chan1,2, Malgorzata Broncel3, Nicole Haseley1, Alice L Herneisen1,2, Emily Shortt1, Moritz Treeck3, Sebastian Lourido1,2* 1 Whitehead Institute for Biomedical Research, Cambridge, MA, USA 2 Biology Department, Massachusetts Institute of Technology, Cambridge, MA, USA 3 Signaling in Apicomplexan Parasites Laboratory, The Francis Crick Institute, London, UK The following chapter is adapted from an article published in eLife (Chan et al., 2023). INTRODUCTION Ca2+-regulated exocytosis is ubiquitous among eukaryotes. This signaling paradigm regulates an array of processes such as neurotransmitter release in neurons, hormone secretion in endocrine cells, and histamine secretion in mast cells (Pang and Südhof 2010). Ca2+-regulated exocytosis is also critical for apicomplexan parasites that are the causative agents of rampant, life-threatening diseases including malaria, toxoplasmosis, and cryptosporidiosis (Havelaar et al. 2015). Central to their pathogenesis is their ability to transition from intracellular replicative stages to extracellular motile stages, which involves a unique form of cellular movement called gliding, egress from the primary host cell, and invasion into a new host. These processes are driven by the Ca2+-regulated exocytosis of apicomplexan-specific membrane-bound organelles called micronemes and rhoptries. The sequential exocytosis of micronemes and rhoptries is required to promote extracellular motile stages of the parasite (V. B. Carruthers and Sibley 1997; I. Blader et al. 2016). Micronemes are localized to the parasite apex and their positioning is dependent on cortical microtubules, ultrastable filaments that polymerize down the length of the parasite (Leung et al. 2017; Chen et al. 2015; Wang et al. 2021). Exocytosis of microneme cargo enables host cell rupture by releasing perforin-like proteins during egress and translocation of exposed adhesins required for gliding and attachment to new host cells (V. B. Carruthers and Sibley 1997; Vern B. Carruthers and Tomley 2008; Kafsack et al. 2009). Multiple microneme proteins, including the associated cysteine 34 repeat modular proteins (CRMP) complex, are also required to trigger the exocytosis of rhoptries upon host cell contact (Kessler et al. 2008; Sparvoli et al. 2022; Sidik et al., n.d.; Singer et al. 2022). Rhoptry proteins include effectors that modulate host responses and transmembrane proteins that are embedded into the host plasma membrane to enable active invasion (M. Lamarque et al. 2011; Tyler and Boothroyd 2011; M. H. Lamarque et al. 2014; Bradley and Sibley 2007; Ong, Reese, and Boothroyd 2010). Intracellular Ca2+ release is necessary and sufficient to trigger the rapid trafficking and exocytosis of micronemes (Sidik, Huet, et al. 2016b; V. B. Carruthers, Giddings, and Sibley 1999; V. B. Carruthers, Moreno, and Sibley 1999; Endo, Sethi, and Piekarski 1982). While Ca2+ is also necessary for rhoptry discharge, exocytosis relies on additional cellular processes such as microneme exocytosis (Coleman et al. 2018; Segev-Zarko et al. 2022). While the exocytosis of micronemes and rhoptries is known to be critical for parasite motility, the mechanisms linking Ca2+ signaling to their trafficking and fusion to the plasma membrane are still unclear. Ca2+ signals in apicomplexans are primarily transduced by Ca2+-dependent protein kinases (CDPKs) (Lourido and Moreno 2015; Billker, Lourido, and Sibley 2009; Farrell et al. 2012; Lourido, Tang, and Sibley 2012; Lourido et al. 2010; Luo, Ruiz, and Moreno 2005; Márquez-Nogueras et al. 2021; Kumar et al. 2017; McCoy et al. 2012; Garrison et al. 2012; Sebastian et al. 2012). CDPKs are serine/threonine protein kinases that are unique to apicomplexans and plants. CDPKs are activated by directly binding to Ca2+, in contrast to Ca2+/calmodulin-dependent protein kinases (CaMKs) found in animals, which are indirectly regulated by Ca2+-bound calmodulin (CaM) (Wernimont et al. 2010; Ojo et al. 2010). CDPKs are crucial for apicomplexan infection—yet are absent from mammals—making them attractive drug targets; however, their mechanisms of action are still not well understood at a cellular and molecular level. In T. gondii, Ca2+- dependent protein kinase 1 (CDPK1) is required for the Ca2+-regulated exocytosis of micronemes, impacting all steps of parasite motility including egress, gliding, and invasion (Lourido et al. 2010). Small-molecule competitive inhibitors against CDPK1 have been identified and have shown some activity against T. gondii (Lourido, Zhang, et al. 2013; Lourido et al. 2010; Doggett et al. 2014; Winzer et al. 2015; Johnson et al. 2012). 35 Identifying the signaling pathways regulated by CDPK1 could reveal the pathways controlling microneme and rhoptry exocytosis. The Ca2+ that activates CDPK1 and other cellular processes is released from intracellular stores following cyclic nucleotide–mediated activation of protein kinase G (PKG) (Sidik, Hortua Triana, et al. 2016; Brown, Long, and Sibley 2017; Bisio and Soldati- Favre 2019). This process can be artificially induced by treating parasites with cGMP specific phosphodiesterase (PDE) inhibitors zaprinast or BIPPO that indirectly activate PKG (Figure 1A)(Lourido, Tang, and Sibley 2012; Yuasa et al. 2005; Sidik, Hortua Triana, et al. 2016; Nofal et al. 2022). PDE inhibition activates PKG within seconds and triggers the Ca2+ and lipid signaling nodes controlling parasite motility, including the exocytosis of micronemes (Figure 1A)(Lourido, Tang, and Sibley 2012; Yuasa et al. 2005). The identity of the CDPK1 substrates that contribute to the regulation of microneme exocytosis remains unknown. Determining CDPK1-dependent phosphorylation is challenging because the signaling pathways controlling parasite motility are rapid and integrate signals from multiple kinases. Global phosphoproteomic studies found that Ca2+-dependent phosphorylation included proteins involved in signal transduction, motility, exocytosis, cytoskeleton, in addition to many proteins with unknown functions; however, the functional relevance and organization of these proteins—especially CDPK1—in regulating exocytosis remain unclear (Treeck et al. 2011; Herneisen et al. 2022; Invergo et al. 2017; Treeck et al. 2014; McCoy et al. 2012; Nofal et al. 2022). Here, we utilized time-resolved phosphoproteomics and chemical genetics to identify 163 proteins phosphorylated by CDPK1. Our comprehensive analysis of CDPK1 not only identified phosphoregulation of known factors involved in parasite motility, but also revealed new regulators of exocytosis. We identified a conserved HOOK complex that is phosphorylated by CDPK1 and is required for microneme exocytosis by mediating the rapid trafficking of micronemes during parasite motility. 36 Figure 1. Identifying CDPK1-dependent phosphorylation with sub-minute resolution. (A) Stimulating parasites with zaprinast triggers Ca2+- mediated activation of CDPK1, resulting in the secretion of microneme proteins (red) required for motile stages of the parasite. Conditional knockdown (cKD) of CDPK1 endogenously tagged with mNeonGreen-mAID-Ty (green) after auxin treatment. (B) Flow cytometry of mNeonGreen (mNG) fluorescence in extracellular CDPK1 cKD or parental TIR1 parasites treated with vehicle or auxin for 3.5 hrs. (C) Schematic of phosphoproteomic time course. Parasites were harvested prior to CDPK1 depletion with auxin for 3.5 hrs, followed by stimulation with zaprinast or vehicle (DMSO). Samples were collected at 0, 9, 30, and 300 sec. The experiment was performed in biological replicates. Samples were labeled with TMTpro, pooled for analysis, and phosphopeptides were enriched using SMOAC prior to LC-MS/MS. Four sets of samples were generated: enriched phosphoproteomes for zaprinast [1] and DMSO [3], and proteomes for zaprinast [2] and DMSO [4]. Mock reporter ion intensities enabling relative quantification for a given peptide are shown to illustrate fold-change of unique phosphopeptide abundances during zaprinast stimulation. CDPK1-dependent phosphorylation is determined by calculating the area under the curve (AUC) difference between vehicle and auxin treatment conditions. (D) Protein abundances in the zaprinast proteome set [2] at 300 sec comparing vehicle- and auxin-treated CDPK1 cKD parasites. (E) UpSet plot for the number of phosphopeptides identified in the enriched zaprinast phosphoproteome [1] across individual replicates. Phosphopeptides exhibiting CDPK1-dependent phosphorylation with p < 0.05 are indicated. (F) Scatter plot of AUC values of enriched zaprinast difference phosphopeptides [1] across biological replicates. Significance was determined by comparing the distribution of AUC values from zaprinast difference phosphopeptides [1] to a null distribution of DMSO phosphopeptides [3] for individual replicates. (G) CDPK1-dependent and zaprinast-dependent phosphopeptide abundances over time. Ratios of zaprinast-treated samples relative to the vehicle-treated (no auxin) t = 0 samples. Median ratios of a group (solid lines). Individual phosphopeptides (opaque lines). CDPK1-dependent phosphopeptides (Group A) determined as described in F. Zaprinast-dependent phosphopeptides (Groups B, C, and D) were determined by comparing the distribution of AUC values from zaprinast vehicle phosphopeptides [1] to a null distribution of DMSO phosphopeptides [3]. Groups were determined by projection-based clustering. (H) GO terms enriched among phosphopeptides undergoing a significant change after zaprinast stimulation. Significance was determined using a hypergeometric test. 37 RESULTS Identifying CDPK1-dependent phosphorylation with sub-minute resolution We examined the effect of CDPK1 on the phosphoproteome. To control the expression of CDPK1, we endogenously tagged the kinase with a C-terminal auxin- inducible degron (AID) for rapid conditional knockdown upon treating parasites with auxin (Figure 1A)(Brown, Long, and Sibley 2018; Smith et al. 2022; Shortt et al. 2022). CDPK1 was robustly depleted from extracellular parasites following auxin treatment for 3.5 hrs (Figure 1B). We compared vehicle- and auxin-treated parasites at four timepoints in the 5 min following zaprinast stimulation (0, 9, 30, and 300 s) to capture the earliest changes following Ca2+ flux (Figure 1C). We used TMTpro labeling to multiplex 16 samples enabling analysis of a complete time course comparing vehicle- and auxin- treated parasites in biological duplicate in a single LC-MS/MS experiment (Li et al. 2020). Samples treated with DMSO instead of zaprinast served as a control. We enriched for phosphopeptides using sequential metal-oxide affinity chromatography (SMOAC) (Tsai et al. 2014). In total, we generated four datasets: an enriched zaprinast phosphoproteome [1], a zaprinast proteome (unenriched) [2], an enriched DMSO phosphoproteome [3], and a DMSO proteome (unenriched) [4]. Of the 4,255 parasite proteins quantified in the zaprinast proteome by LC-MS/MS, CDPK1 was the only protein with a greater than two-fold depletion in parasites treated with auxin (Figure 1D). The remainder of the proteome was largely stable (median log2-fold change = -0.002 ± 0.089 M.A.D). We first quantified peptide abundances across time relative to the 0 sec vehicle- treated time point to identify CDPK1-dependent phosphorylation, which yielded kinetic profiles of individual phosphopeptides in vehicle and auxin conditions (Figure 1C). The zaprinast phosphoproteome included 2,570 phosphorylated proteins, represented by 10,594 unique phosphopeptides quantified across both biological replicates (Figure 1E). We calculated the area under the curve (AUC) of individual phosphopeptide profiles for vehicle (AUCvehicle) and auxin (AUCauxin) conditions. To identify the subset of phosphopeptides exhibiting CDPK1-dependent phosphorylation, we calculated the 38 difference between AUC values (AUCdifference) by comparing the distribution of AUCdifference values in the zaprinast phosphoproteome to a null distribution derived from the DMSO phosphoproteome (Figure 2A). This approach allowed us to account for the variability among AUCs under conditions that did not exhibit dynamic changes. We identified 74 unique CDPK1-dependent phosphopeptides across both biological replicates (Group A), belonging to 69 proteins (Figure 1E–F). Additionally, we identified peptides that were zaprinast responsive—despite being CDPK1 independent—by comparing the distribution of AUCvehicle values in the zaprinast phosphoproteome to a null distribution derived from the DMSO phosphoproteome, resulting in 809 unique phosphopeptides representing 501 proteins. We utilized projection-based clustering to sort kinetic profiles of CDPK1-independent peptides into three groups, in which Group B and Group C contained phosphopeptides that increased in abundance and Group D contained phosphopeptides that decreased in abundance (Thrun and Ultsch 2021)(Figure 1G, Figure 2B) (Thrun and Ultsch 2021). Significant changes at the peptide level are attributed to altering levels of phosphorylation as opposed to protein levels. Of the 543 proteins exhibiting zaprinast-dependent phosphorylation, 484 were quantified at the proteome level. When comparing the 300 to 0 sec time points, zaprinast-dependent proteins displayed no significant changes (median log2-fold change = 0 ± 0.074 M.A.D). This was consistent with the rapid time scale of these experiments and the overall stability of the 4,255 proteins quantified in the proteome (median log2-fold change = 0.01 ± 0.089 M.A.D). Gene ontology analysis identified several classes of genes that may be relevant to the regulation of zaprinast responses within the different groups (Figure 1H). CDPK1- independent phosphorylation and dephosphorylation was prevalent amongst proteins regulating cyclic nucleotide signaling (ACβ, GC, UGO, PDE1, PDE2, PDE7, PDE9, PDE10, and PKG), lipid signaling (PI4K, PI4P5K, PI-PLC, DGK1, and PAP1), and Ca2+ signaling (CDPK2A and CDPK3). CDPK1-independent phosphorylation was also observed on downstream proteins regulating microneme exocytosis (APH and DOC2.1), rhoptry exocytosis (ARO, AIP, PL3, and NdP2), gliding motility (AKMT and MyoA), and homeostasis/biogenesis (NHE3, VHA1, NST2, and ERK7). MyoA phosphorylation was 39 observed at S20 and S21, which was previously shown to be phosphorylated by CDPK3 and required for gliding motility (Gaji et al. 2015). SCE1 phosphorylation was observed at S225, which was previously shown to be phosphorylated by CDPK3 and required to relieve inhibition during Ca2+-stimulated egress (McCoy et al. 2017). Our analysis highlights temporally-resolved changes in the phosphoproteome in response to zaprinast stimulation. We identify a subset of proteins phosphorylated by CDPK1 that likely represent factors involved in motility-related exocytosis, which we discuss in further detail below. Figure 2. Zaprinast-dependent phosphoproteome. (A) Distribution of AUCdifference values for individual peptides in the enriched DMSO phosphoproteome and enriched zaprinast phosphoproteome across two biological replicates. (B) Heatmap of zaprinast-dependent phosphopeptide abundance ratios across time relative to the vehicle t0 interval during auxin or vehicle treatment. Peptides are organized by CDPK1- dependent phosphopeptides (Group A) and CDPK1-independent phosphopeptides (Group B, C, D). 40 Myristoylation modulates CDPK1 activity and alters its interacting partners CDPK1 was found to be myristoylated on Gly2 (Broncel et al. 2020). Myristoylation results in lipid modifications on proteins that can impact membrane targeting and protein-protein interactions (Martin, Beauchamp, and Berthiaume 2011; Wright et al. 2014). The myristoylated proteome of T. gondii was recently characterized using metabolic tagging and enrichment (Broncel et al. 2020). In this approach, parasites were grown in the presence of a myristic acid analogue containing a terminal alkyne group (YnMyr) that allowed enrichment of labeled proteins through click chemistry (Heal et al. 2011). We assessed whether myristoylation of CDPK1 affects the kinase’s function. We validated myristoylation of CDPK1 using myristoylation-dependent pull downs followed by immunoblot detection and MS analysis (Figure 3A–B). To investigate the role of myristoylation on CDPK1 function, an inducible knock-down strain (iKD) was generated by introducing a mAID-Myc tag at its C terminus (Figure 3C–D). Auxin-dependent depletion was confirmed using immunoblot (Figure 3D). Depletion of CDPK1 abolished ionophore-induced egress, as expected from previous results (Figure 3E)(Lourido et al. 2010). Next, we complemented the iKD parasites by introducing HA-tagged WT (cWT) or myristoylation defective (cMut, G2A) copies of CDPK1 into the UPRT locus (Figure 4A). We verified correct integration of both complementation constructs (Figure 3E), and confirmed their equivalent and constitutive expression, as well as the auxin sensitivity of the endogenous mAID-tagged CDPK1 (Figure 4B). As predicted, the cMut allele was not myristoylated, based on acylation-dependent pull downs and immunoblotting (Figure 4C). 41 Figure 3. CDPK1 myristoylation, inducible knockdown, and complementation. (A) YnMyr-dependent pull down confirming myristoylation of CDPK1. GRA2 antibody was used as a loading control. (B) MS2 fragmentation spectra indicating myristoylation of Gly2 of CDPK1 after YnMyr-dependent pull down. (C) PCR analysis confirming correct integration of the mAID cassette at the C terminus of endogenous CDPK1 in the TIR1 line. (D) Immunoblot validation of auxin-dependent depletion of CDPK1 in the iKD line using the anti-Myc antibody and the anti-toxo antibody as a loading control. The band at 75 kDa represents anti-Myc-related background. (E) Conditional depletion of CDPK1 abolishes ionophore-induced egress from host cells. Intracellular parasites were treated with auxin or vehicle (EtOH) for 2 hrs and egress was initiated by addition of 8 µM A23187. The number of intact vacuoles was monitored over the course of 6 min. Each data point is an average of two biological replicates, each in technical triplicate, error bars represent standard deviation. (F) PCR analysis confirming correct integration of the complementation constructs encoding the WT (cWT) and myristoylation mutant (cMut) copies of CDPK1 at the UPRT locus of the iKD line. Primers are indicated by arrows. Base pairs (bp). (G) In the absence of auxin, both cWT and cMut parasites egress from host cells within 2 min post-ionophore treatment. 42 Given that myristoylation is frequently reported to facilitate membrane association, we examined the localization of cWT and cMut CDPK1 by immunofluorescence (Figure 4D). No clear differences were detected between the punctate cytosolic patterns of cWT and cMut. We explored possible effects of myristoylation on the subcellular fractionation of CDPK1 resolved using differential centrifugation (Figure 4E). First, we evaluated the fractionation pattern of the endogenous, myristoylated CDPK1. YFP-expressing parasites were metabolically labeled with Myr or YnMyr and lysed in a hypotonic buffer to preserve intact membrane structures. Next, lysates were fractionated to generate a low-speed pellet and supernatant at 16,000 x g. The low-speed supernatant was fractionated further into a high-speed pellet and supernatant at 100,000 x g. Click reaction based pull down and immunoblotting were used to resolve myristoylation-dependent partitioning. In contrast to the doubly acylated GAP45, which was present exclusively in the low-speed pellet, myristoylated CDPK1 was observed in the low-speed pellet and supernatant, and the high-speed pellet (Figure 4E), suggesting a potential association with membranous structures or higher molecular weight complexes. We next used both the cWT and cMut lines to elucidate any myristoylation-dependent changes to CDPK1 localization. While cWT could be found predominantly in the high-speed pellet, loss of myristoylation released cMut into the high-speed supernatant confirming an association with membranous structures or large protein complexes (Figure 4F). To evaluate the role of CDPK1 myristoylation in parasite fitness, we performed plaque assays comparing the various strains (Figure 4G). In the presence of the endogenous copy of CDPK1, both complemented lines developed normally. However, upon auxin-mediated depletion of endogenous CDPK1, cMut plaque size substantially decreased. This finding demonstrates that one or more steps of the T. gondii lytic cycle are negatively affected by the loss of CDPK1 myristoylation. In light of CDPK1’s known function, we next explored whether CDPK1 myristoylation might impact the parasite’s ability to egress from host cells. In the absence of auxin, cWT, cMut, and the parental line (iKD) egressed within two min of ionophore stimulation (Figure 3G). While cWT parasites maintained similar egress kinetics following auxin treatment, cMut parasites 43 showed a significant delay after two min of treatment (Figure 4H). This egress delay was overcome by six min, suggesting that CDPK1 myristoylation is important for ionophore- induced egress, but not essential. Figure 4. Myristoylation modulates CDPK1 activity and alters its interacting partners. (A) Complementation strategy used to evaluate the functional importance of CDPK1 myristoylation. See Figure 2—figure supplement 1 for the construction of the iKD line. (B) Immunoblot demonstrating the auxin-dependent depletion of endogenous CDPK1 in the iKD, cWT, and cMut parasites (Myc) as well as equivalent expression of the complements (HA). T. gondii (toxo) antibody was used as a loading control. (C) Biochemical validation of complemented lines by YnMyr-dependent pull down. Enrichment of WT and Mut complements (HA). The inducible endogenous CDPK1 (Myc) and T. gondii (toxo) antibody was used as enrichment and loading controls, respectively. (D) Localization of the complemented versions of CDPK1 and corresponding cytosolic reporters within cWT (GFP) and cMut (mCherry) by immunofluorescence. (E) Myristoylation-dependent subcellular partitioning of CDPK1. Localization of YnMyr-enriched CDPK1 was evaluated using differential centrifugation. The partitioning into pellet [P] and supernatant [S] fractions was detected by immunoblot (CDPK1) and compared to doubly acylated GAP45. GFP and SAG1 were used as S and P controls, respectively. As only half of the supernatant fraction was removed from the high-speed pellet (100,000 x g), the GFP signal is present in the latter. (F) Partitioning of complemented WT and mutant CDPK1 after high speed centrifugation (HA). T. gondii (toxo) antibody was used as a P control whereas GFP and mCherry were used as S controls for cWT and cMut, respectively. (G) Plaque assays demonstrating that myristoylation of CDPK1 is important for the lytic cycle of T. gondii. (H) Lack of CDPK1 myristoylation delays ionophore-induced egress from host cells. Each data point is an average of n = 3 biological replicates, error bars represent standard deviation. Significance calculated using 1-way ANOVA with Tukey’s multiple comparison test. See Figure 2—figure supplement 1 for vehicle controls. (I) Immunoprecipitation-MS (IP- MS) of CDPK1-HA in cWT, cMut, and untagged TIR1 parasites across n = 2 biological replicates. Significantly enriched proteins (red) for proteins with more than three unique peptides, pcWT < 0.05, and pcMut< 0.05, and log2 fold-change > 1 across both pull-downs; t-tests and Benjamini-Hochberg corrected. (J) IP-MS fold-enrichment comparing cWT and cMut pull-downs. Significantly enriched proteins (red) for proteins with more than 3 unique peptides, p < 0.05, and log2 fold-change > 1 or < -1; t-test and Benjamini-Hochberg corrected. 44 Finally, we wanted to examine whether myristoylation of CDPK1 affects its interactions with other proteins, which may occur by direct binding to the kinase or by indirect association with a shared membrane structure. We performed immunoprecipitation mass spectrometry (IP-MS) on HA-tagged CDPK1 from cWT, cMut, and untagged TIR1 parasites after hypotonic lysis. CDPK1 was the most significantly enriched protein across both cWT and cMut pulldowns when compared to untagged controls (Figure 4I). Two additional proteins were significantly enriched along with CDPK1: NUP134 (TGGT1_240510) and SRS36D (TGGT1_292280). Comparing the cWT and cMut pulldowns, we found two proteins preferentially associated with cWT: NUDIX hydroxylase (TGGT1_227450) and a 14-3-3 protein (TGGT1_269960) (Figure 4J). Loss of myristoylation appears to enhance interactions with three different proteins: ROM4 (TGGT1_268590), a putative T complex protein 1 alpha subunit (TGGT1_229990), and IMP dehydrogenase (TGGT1_233110). Genome-wide knockout screen data suggest that NUP134 and the putative T complex protein 1 are the only putative interacting partners required for parasite fitness (Sidik, Huet, et al. 2016b). NUP134 binds and co-localizes with the nuclear pore complex, whereas the putative T complex protein 1 may function as an ATP-dependent chaperone (Courjol et al. 2017). CDPK1 is localized primarily to the cytoplasm of the parasite and partially in the nucleus, which could explain its interactions with NUP134 (Ojo et al. 2010; Lourido et al. 2010). ROM4 is an integral membrane protease required to shed secreted microneme proteins through proteolysis, and its knockdown impaired gliding motility and invasion of host cells (Buguliskis et al. 2010). ROM4 localizes to the plasma membrane of parasites but was enriched with cytosolic CDPK1 and was not observed in our sub-minute resolution phosphoproteomics. Our pulldowns did not reveal a membrane compartment association for myristoylated CDPK1 but did identify interacting proteins enriched in cytosolic and myristoylated CDPK1. Given the annotated functions of these interacting proteins, the factors are unlikely to participate in the motility-related functions of CDPK1. Additional studies will be required to understand how myristoylation influences CDPK1 activity. 45 Identification of direct CDPK1 targets through thiophosphorylation Our phosphoproteomic time course experiment identified proteins exhibiting CDPK1-dependent phosphorylation in live parasites, which includes direct and indirect substrates of the kinase. CDPK1 is unusual among apicomplexan and metazoan kinases in that it contains a glycine at its gatekeeper residue, resulting in an expanded ATP- binding pocket that can accommodate bulky ATP analogues. CDPK1 can use bulky analogues like N6-furfuryladenosine (kinetin)-5′-O-[3-thiotriphosphate] (KTPγS) to thiophosphorylate its substrates in parasite lysates (Lourido, Jeschke, et al. 2013). Thiophosphorylated substrates can subsequently be enriched with an iodoacetyl resin (Blethrow et al. 2008). Specificity can be assessed by comparison to mutant parasites harboring a G128M gatekeeper mutation in CDPK1 (CDPK1M) that retains kinase activity but prevents it from using KTPγS. Previous studies using lysates identified a small set of six putative CDPK1-dependent substrates; however, these studies lost the target specificity conferred by the subcellular context and lacked the accuracy and sensitivity of quantitative proteomic approaches (Lourido, Jeschke, et al. 2013). We implemented several modifications to the thiophosphorylation procedure that improved identification of CDPK1 targets (Figure 5A)(Rothenberg et al. 2016). First, we used stable isotope labeling of amino acids in cell culture (SILAC) to directly compare thiophosphorylation in WT (CDPK1G) and mutant (CDPK1M) parasites. Second, we used parasites semi-permeabilized with the bacterial toxin aerolysin—as opposed to the preparations of detergent-lysed parasites used in prior methods. Aerolysin forms 3-nm pores in the plasma membrane which permit the diffusion of small molecules but not proteins, enabling us to perform labeling reactions without drastically disrupting the concentration or localization of proteins (Iacovache et al. 2006). Lastly, we prevented non-specific extracellular substrate labeling due to premature lysis by treating semi- 46 Figure 5. Identifying the direct substrates of CDPK1. (A) Schematic describing a strategy to identify direct substrates of CDPK1. WT (CDPK1G) and mutant (G128M; CDPK1M) parasites were grown in SILAC media for multiplexed quantitation. Extracellular parasites were semi-permeabilized with aerolysin, enabling diffusion of small molecules but not proteins. CDPK1 substrate labeling was initiated by treating semi-permeabilized parasites with Ca2+, KTPγS, ATP, and 1B7. While CDPK1 in both WT and mutant parasites can utilize ATP to phosphorylate substrates, only WT parasites can use KTPγS to thiophosphorylate substrates. Thiophosphorylated peptides were specifically enriched and the remaining flow-through was saved for whole proteome analysis. Enriched and whole proteome samples were analyzed by LC-MS/MS. (B) 1B7 nanobody treatment inhibits non-specific extracellular kinase activity of CDPK1. Thiophosphorylated substrates were detected in lysates using an anti-thiophosphate ester antibody immunoblot. Extracellular CDPK1 activity (lane 1) was blocked by 1B7 (lane 2). Aerolysin treatment resulted in intracellular labeling (lane 5) that was unaffected by 1B7 (lane 6). (C) Thiophosphorylation performed in aerolysin-treated parasites comparing WT (CDPK1G) and mutant (CDPK1M) strains. Detection was performed as in B. Tubulin was used as a loading control. (D) Heatmap quantification of peptides using LC-MS/MS. Fold-change of peptide abundance shown as a ratio of WT (CDPK1G) to mutant (CDPK1M) abundances. Experiment was performed in n = 3 biological replicates. (E) Abundances of unique peptides after thiophosphorylation in CDPK1G and CDPK1M parasites across n = 3 biological replicates. Significantly enriched phosphorylated peptides are colored in red (-log (p)*fold-change > 4), one-tailed t-test. (F) GO terms enriched among significant phosphopeptides from E. Significance 10 was determined using a hypergeometric test. (G) Putative targets of CDPK1 determined by sub-minute phosphoproteomics and thiophosphorylation of direct substrates. For a given CDPK1 target gene, the presence of a unique peptide phosphorylated in a CDPK1- dependent manner (column 1) is indicated if identified in the time course (green) and/or thiophosphorylation (magenta). The presence of additional unique phosphorylated peptides exhibiting zaprinast-dependent effects (column 2) is indicated if the peptide was phosphorylated (orange) or dephosphorylated (blue). Numbered boxes indicate multiple unique peptides. Fitness scores (column 3) obtained from genome- wide KO screen data (blues). Lower scores indicate gene is required for lytic stages of the parasite. Gene names (left), TGGT1 gene IDs (right). Gene names with asterisks (*) are associated with published data. (H) Signaling diagram describing parasite motility. Proteins exhibiting CDPK1-dependent phosphorylation by either sub-minute phosphoproteomics or thiophosphorylation are indicated (green). Proteins exhibiting CDPK1-independent phosphorylation (red) or dephosphorylation (blue) are indicated. 47 permeabilized parasites with 1B7, a nanobody that allosterically inhibits CDPK1 but does not enter the cytosol of semi-permeabilized parasites (Figure 5B)(Ingram et al. 2015). Thiophosphorylation experiments were performed in biological triplicate comparing CDPK1G and CDPK1M parasites. We assessed thiophosphorylation labeling by immunoblot, identifying an array of proteins specifically labeled in CDPK1G, but not CDPK1M parasites (Figure 5C). MS analyses quantified the abundance of peptides in CDPK1G relative to CDPK1M parasites, identifying 734 unique peptides across three biological replicates (Figure 5D). Samples from CDPK1G parasites were significantly enriched in phosphorylated peptides, consistent with CDPK1-mediated thiophosphorylation of targets (Figure 5E). 123 peptides across 104 proteins were likely direct substrates of CDPK1. GO enrichment analysis did not reveal any pathways relevant to the function of CDPK1 in exocytosis (Figure 5F). While our approach largely maintains kinases in their subcellular context, aerolysin treatment disrupts native ion concentrations and detaches the plasma membrane from the inner membrane complex (IMC) (Wichroski et al. 2002). We therefore proceeded to consider the thiophosphorylated substrates in the context of the time-resolved phosphoproteomics. CDPK1 targets participate in pathways controlling parasite motility Considering the thiophosphorylation and time-resolved phosphoproteomics, we arrived at a prioritization scheme to identify 163 proteins phosphorylated in a CDPK1- dependent manner (Figure 5G). Proteins were divided into five classes based on the overlap of phosphorylated sites between both approaches. Class 1 contains five proteins for which the same phosphorylated site was identified in both the time course and thiophosphorylation experiments. Class 2 contains four proteins for which phosphorylated sites identified across both approaches were within 50 amino acid residues of one another. Class 3 contains two proteins that were enriched by thiophosphorylation and were also CDPK1-dependent in the time course, but the identified sites were more than 50 residues apart. Class 4 contains 93 proteins that were exclusively enriched by thiophosphorylation. Lastly, Class 5 contains 59 proteins that were CDPK1-dependent exclusively in the time course and are likely indirect targets of 48 the kinase. Proteins within each class were further stratified by fitness scores reported from genome-wide knockout screens, with lower scores representing genes more essential for parasite fitness in cell culture (Sidik, Huet, et al. 2016b). Of the 163 targets of CDPK1, 72 proteins across all classes also displayed changes in zaprinast-dependent phosphorylation at distinct sites that were independent of CDPK1. This overlap suggests that some proteins modified by CDPK1 are also regulated by additional kinases and phosphatases. We hypothesized that regulators of exocytosis would be found among the targets of CDPK1. Of the 163 protein targets, 38 have been previously localized and/or functionally characterized (indicated with asterisks; Figure 5G). Of these 38 proteins, 13 have been implicated in regulating motile stages of the parasite (Figure 5H, Figure 6A). Among the candidates associated with microneme exocytosis, centrin 2 (CEN2) and RNG2 have previously been localized to tubulin-based structures in the apical complex of the parasite, and their knockdown is sufficient to inhibit secretion of microneme proteins and block host cell invasion (Leung et al. 2019; Lentini et al. 2019; Katris et al. 2014). Candidates associated with rhoptry exocytosis include the armadillo repeats only protein (ARO), ARO-interacting protein (AIP), palmitoyl acyltransferase DHHC7, and patatin-like phospholipase (PL3) (Beck et al. 2013; Mueller et al. 2013, 2016; Wilson et al. 2020). ARO and DHHC7 both localize to the cytosolic face of rhoptries, where they influence the recruitment of AIP and ACꞵ. Knockdown of either DHHC7 or ARO disrupts the localization of mature rhoptries, inhibiting the secretion of rhoptry proteins required for invasion. Another potential CDPK1 target, the apical complex lysine methyltransferase (AKMT), rapidly relocalizes from the apical complex to the parasite body during Ca2+-regulated motility, and its knockdown impairs the gliding motility of parasites required for invasion and egress (Heaslip et al. 2011). Other putative CDPK1 targets participate in homeostasis and biogenesis of secretory organelles, such as the vacuolar type Na+/H+ exchanger (NHE3), dynamin-related protein B (DrpB), V-ATPase a1 (VHA1), and GDP-fucose transporter (NST2). NHE3-knockout parasites exhibit sensitivity to osmotic shock and dysregulated cytosolic Ca2+, resulting in reduced secretion of microneme proteins and an inhibition of invasion (Francia et al. 2011). DrpB was 49 previously identified as a CDPK1 target, and its depletion results in severe defects in the biogenesis of micronemes and rhoptries (Breinich et al. 2009; Lourido, Jeschke, et al. 2013). VHA1 and NST2 participate in the maturation of microneme and rhoptry proteins (Stasic et al. 2019; Bandini et al. 2019). Lastly, signaling proteins regulating intracellular cGMP levels, such as the guanylate cyclase (GC) and the unique GC organizer (UGO), were also phosphorylated in a CDPK1-dependent manner (Brown and Sibley 2018; Yang et al. 2019; Bisio et al. 2019). CDPK1 acts downstream of GC-mediated production of cGMP, which may suggest regulatory feedback on Ca2+ release, as has been suggested for CDPK3 (Nofal et al. 2022). These data demonstrate that identification of CDPK1 targets can uncover proteins involved in Ca2+-regulated exocytosis. Figure 6. Factors controlling parasite motility. (A) Expanded list of factors involved in parasite motile stages from Fig. 3H. DISCUSSION In this study, we sought to identify new regulators of Ca2+-mediated exocytosis in T. gondii by studying the targets of a key regulator, the kinase CDPK1. We identified 163 proteins phosphorylated in a CDPK1-dependent manner using sub-minute resolution phosphoproteomics and thiophosphorylation for direct substrate capture. We determined that myristoylation of CDPK1 contributes to the kinase’s function during the lytic cycle. 13 of the identified CDPK1 targets have previously described functions in 50 parasite motility, revealing possible points of regulation within relevant signaling pathways. Over the past decade, several efforts have sought to characterize phosphorylation within apicomplexans. Improvements in phosphopeptide enrichment and mass spectrometry have enabled global characterization of the phosphoproteomes from T. gondii tachyzoites and P. falciparum schizonts, trophozoites, and ring-stages(Treeck et al. 2011; Solyakov et al. 2011; Lasonder et al. 2012; Pease et al. 2013). Coupling such approaches with genetic and pharmacological tools has enabled the deconvolution of kinase-specific effects from the vast global phosphorylation program(Treeck et al. 2014; Nofal et al. 2022; Fang et al. 2017; Invergo et al. 2017; Pease et al. 2018; Blomqvist et al. 2020), revealing the activation of MyoA-mediated gliding by TgCDPK3, the roles of PbCDPK4 and PbSRPK1 during male gametogenesis, and the erythrocytic-stage effects of PfCDPK1, PfPK7, and PfCDPK5(Tang et al. 2014; Gaji et al. 2015; Invergo et al. 2017; Fang et al. 2017; Pease et al. 2018; Blomqvist et al. 2020). Attributing individual phosphorylation events to specific kinases can be complicated by cumulative changes or adaptation of the signaling networks in knockouts. Chemical-genetic approaches can more precisely inhibit the activity of a given kinase at the time of the assay, while controlling for off-target effects, and have been used to characterize PfPKG-dependent phosphorylation (Alam et al. 2015; Brochet et al. 2014). Kinetically-resolved phosphoproteomics has enabled a more nuanced understanding of signaling cascades during parasite motility(Invergo et al. 2017; Herneisen et al. 2022; Nofal et al. 2022). While such technical advances have led to more in-depth phosphoproteome mapping, identifying the direct targets of a given kinase remains a key challenge to structuring the observed changes into signaling pathways. To resolve direct and indirect effects on the phosphoproteome for T. gondii CDPK1, we combined two complementary approaches: temporally-resolved phosphoproteomics and bio-orthogonal labeling of direct substrates. We implemented several methodological advances. Rapid conditional knockdown using the auxin- inducible degron system resulted in rapid and specific depletion of the kinase of interest for the phosphoproteomic studies(Brown, Long, and Sibley 2018). Additionally, 51 quantification and coverage by mass spectrometry were improved through TMTpro multiplexing and ion mobility spectrometry (FAIMS)(Van Vranken, Pontano Vaites, and Schweppe, n.d.; Bekker-Jensen et al. 2020). We also improved on the low coverage observed in previous thiophosphorylation experiments with parasite lysates(Lourido, Jeschke, et al. 2013; Fang et al. 2017) by maintaining a more native signaling environment using aerolysin semi-permeabilization, SILAC-based peptide quantification, and shortened labeling times(Rothenberg et al. 2016). These approaches characterize proteome-wide phosphorylation kinetics in live parasites and reveal direct kinase- substrate relationships for CDPK1. Myristoylation of CDPK1 contributes to its function. Contrary to previous reports describing the kinase as cytosolic or nuclear(Ojo et al. 2010; Pomel, Luk, and Beckers 2008), we demonstrate that myristoylated CDPK1 is at least partially associated with structures that can be fractionated from the cytoplasm by differential centrifugation. Loss of myristoylation led to CDPK1 release from the insoluble fraction. The ortholog of CDPK1 in P. berghei, PbCDPK4, also displays myristoylation-specific functions during male gametogenesis: myristoylated PbCDPK4 is critical for the first genome replication, whereas the non-myristoylated PbCDPK4 is important for the completion of gametogenesis(Fang et al. 2017). Myristoylation could impact CDPK1’s ability to access certain targets efficiently. However, such interactions appear to be too weak or transient to be captured by immunoprecipitation. Mutating the myristoylation site of CDPK1 only modestly affected ionophore-induced egress. However, minor effects in these key transitions may be magnified over repeated lytic cycles, which may explain the more substantial impact of losing CDPK1 myristoylation on plaque formation. Alternatively, other kinases may compensate for decreased CDPK1 activity, particularly under hyperactivated conditions like stimulated egress. A plausible candidate for compensation is CDPK2A, which also appears to be myristoylated, despite lacking an N-terminal MG motif, and was recently shown to display epistatic interactions with CDPK1(Broncel et al. 2020; Shortt et al. 2022). CDPK1 plays a critical role in the transition from the replicative intracellular stages to motile extracellular stages (Lourido et al. 2010; Shortt et al. 2022). During this 52 transition parasites must execute rapid cellular changes that involve the exocytosis of micronemes and rhoptries, reorganization of the cytoskeleton, gliding motility, and maintenance of ion homeostasis(I. J. Blader et al. 2015). CDPK1 has also been suggested to control the actomyosin system and extrusion of the conoid(Tosetti et al. 2019). Our detailed target analysis suggests CDPK1 regulates—and possibly helps coordinate—multiple pathways. We revealed over a hundred CDPK1 targets with diverse predicted functions, including many relevant to phenotypes dependent on CDPK1. In addition to the effects on microneme trafficking examined in this study, our results point to a direct link between CDPK1 and rhoptry exocytosis. Ca2+ has been implicated in rhoptry discharge, as the rhoptry-localized Ca2+-binding FER2 is required for rhoptry exocytosis(Coleman et al. 2018). We observed a preponderance of CDPK1-dependent phosphorylation on proteins regulating rhoptry exocytosis, including DHHC7, ARO, and AIP. Phosphorylation on ARO was observed near the N-terminal acylation sites required for rhoptry targeting and may regulate the bundling and positioning of mature rhoptries during motile stages(Mueller et al. 2016). However, formally demonstrating the relationship between Ca2+/CDPK1 and rhoptry discharge is complicated by the dependency of the latter on the secretion of certain microneme proteins (V. B. Carruthers and Sibley 1997; Ben Chaabene, Lentini, and Soldati-Favre 2021). T. gondii tachyzoites have several rhoptries, yet only two are docked for exocytosis at a given time (Mageswaran et al. 2021; Aquilini et al. 2021; Segev-Zarko et al. 2022). Regulating the activity of ARO during motile stages may influence the ability to mobilize and re-dock rhoptries in preparation for invasion. Considering how different CDPK1 substrates function, we expect the various phenotypes associated with CDPK1 will depend on distinct sets of substrates. 53 MATERIALS & METHODS Cell culture T. gondii parasites were grown in human foreskin fibroblasts (HFFs, ATCC SRC-1041) maintained in DMEM (GIBCO 11965118) supplemented with 3% inactivated fetal calf serum and 10 µg/mL gentamicin (Thermo Fisher Scientific), referred to as media. When noted, DMEM was supplemented with 10% inactivated fetal bovine serum (IFS) and 10 µg/mL gentamicin, referred to as 10% IFS media. Parasites and HFFs were grown at 37°C/5% CO2 unless indicated otherwise. Parasite transfection and strain construction Genetic background of parasite strains T. gondii RH strains were used as genetic backgrounds for this study. All strains contain the ∆ku80∆hxgprt mutations to facilitate homologous recombination (Huynh and Carruthers 2009). TIR1 expresses the TIR1-FLAG ubiquitin ligase and the CAT enzyme conferring chloramphenicol resistance (Brown, Long, and Sibley 2017). Transfection Parasites were pelleted at 1000 x g for 5 to 10 min and resuspended with Cytomix (10 mM KPO4, 120 mM KCl, 0.15 mM CaCl2, 5 mM MgCl2, 25 mM HEPES, 2 mM EDTA, 2 mM ATP, and 5 mM glutathione) and combined with DNA to a final volume of 400 µL. Parasites were electroporated using an ECM 830 Square Wave electroporator (BTX) in 4 mm cuvettes with the following setting: 1.7 kV, 2 pulses, 176 µs pulse length, and 100 msec interval. Endogenous tagging of CDPK1 (CDPK1-AID) CDPK1-AID was generated in the study (Shortt et al. 2022). Briefly, V5-mNG-mAID-Ty was PCR amplified from pBM050 (V5-TEV-mNG-mAID-Ty; GenBank: OM640006) to attach homology arms to TGGT1_301440 (CDPK1). DNA was co-transfected with a Cas9 expression plasmid targeting CDPK1 in TIR1 parasites. mNG-expressing parasites were isolated using FACS and subcloned by limiting dilution. Tagging was confirmed by PCR, flow cytometry, and immunoblotting. Endogenous tagging of CDPK1 (iKD) and complementation (cWT and cMut) Generating the inducible CDPK1 knock-down strain (iKD CDPK1). The pTUB1_YFP_mAID_3HA vector was amplified by inverse PCR using primer pair P1/P2 to substitute the 3xHA tag sequence for a Myc tag encoding sequence(Brown, Long, and Sibley 2017). A Cas9 expression plasmid targeting the 3’UTR of CDPK1 was generated by inverse PCR on pSag1_Cas9-U6_sgUPRT using primers P3/P4(Shen, Brown, et al. 2014). The sequence encoding mAID-Myc-HXGPRT was PCR amplified using primer pair P5/P6 and co-transfected into TIR1 parasites with the Cas9 expression plasmid. Recombinant parasites were selected 24 hrs post transfection by addition of mycophenolic acid (MPA; 25 µg/mL) and xanthine (XAN; 50 µg/mL) to culture medium. Lines were cloned, and successful 5’ and 3’ integration of the mAID-Myc-HXGPRT 54 cassette was confirmed using primer pairs P30/P31 and P32/P33. Absence of WT was confirmed using primers P34/P35. Generating the cWT and cMut CDPK1 complementation strain. To generate the complementation construct, pUPRT_CDPK1_ HA_T2A_GFP (GenBank: Pending), the CDPK1 5’UTR was amplified from genomic DNA using primer pair P7/P8. Recodonized CDPK1 cDNA-HA sequence was synthesized (GeneArt strings, Life Technologies) and amplified with appropriate overhangs using primers P9/P10. Sequence encoding T2A- GFP was amplified from an in-house unpublished plasmid using primer pair P11/P12. The three resulting fragments were Gibson cloned into the PacI-linearized pUPRT_HA vector(Reese et al. 2011). To generate the complementation construct, pUPRT_CDPK1(G2A)_HA_T2A_mCherry (GenBank: Pending), the CDPK1 5’UTR was amplified from genomic DNA using primer pair P13/P8. Recodonized CDPK1-HA was amplified from pUPRT_CDPK1_ HA_T2A_GFP with appropriate overhangs using primers P14/P15. Primers P13/P15 were used to introduce a G2A point mutation within CDPK1. Sequence encoding T2A-mCherry was amplified from an in-house unpublished plasmid using primer pair P11/P16. The three resulting PCR amplicons were Gibson cloned into the PacI-linearized pUPRT_HA vector. Complementation plasmids were linearized with AclI and individually co-transfected with the Cas9 expression plasmid targeting the UPRT locus. Transgenic parasites were subjected to 5’-fluo-2’-deoxyuridine (FUDR) selection (5 µM) 24 hrs post transfection. Resistant parasites were cloned, and successful 5’ and 3’ integration was confirmed using primer pairs P36/P37 and P38/P39, respectively. Disruption of the endogenous UPRT locus was confirmed using primer pair P40/P41. Sub-minute phosphoproteomics Parasite harvest and treatment T. gondii tachyzoites from the RH strain CDPK1-AID were used to infect nine 15 cm dishes. 3.75 x 107 parasites were used to infect each dish in 20 mL of media. Approximately 24 hrs later, the media of eight dishes was replaced with 15 mL of media containing 1 µM of compound 1 to block egress and synchronize parasites by inhibiting PKG (Hopp, Bowyer, and Baker 2012; R. G. K. Donald et al. 2006; H. M. Taylor et al. 2010). The media of the ninth dish was replaced with media containing DMSO. On day 2, parasites were harvested when the monolayer of HFFs in the DMSO dish were approximately 80% lysed. The eight dishes treated with compound 1 were washed once with 10 mL of warm 1X PBS, once with 30 mL of warm 1X PBS, and incubated with 10 mL of warm FluoroBrite (Fluorobrite DMEM A1896701, 4 mM glutamine, 10 µg/mL gentamicin) for 10 min at 37°C. Infected HFFs were scraped and passed through a 27G needle to mechanically liberate parasites and passed through a 5 µm filter. Parasites were pelleted at 1000 x g at 4°C for 10 min, resuspended in 10 mL of FluoroBrite and a 1:250 dilution was used to count parasites (approximately 2.25 x 109 total parasites). Parasites were pelleted at 1000 x g at 4°C for 7 min and resuspended in 2 mL of FluoroBrite. 1 mL of parasites were diluted into a total volume of 40 mL of FluoroBrite containing either 500 µM of auxin or vehicle of 1X PBS and incubated at 37°C for 3.5 55 hrs. 50 µL of auxin- or vehicle-treated parasites and untagged TIR1 parasites were analyzed by flow cytometry (Miltenyi MACSQuant VYB) to detect mNeonGreen fluorescence. After confirming depletion of CDPK1, parasites were pelleted at 1000 x g for 10 min at room temperature and resuspended in 210 µL of 500 µM auxin or vehicle of PBS diluted in FluoroBrite. To obtain a time course, the 0 sec time point was first collected by mixing 16 µL of parasites with 4 µL of 5X DMSO (0.5% DMSO in FluoroBrite) and immediately lysed with 20 µL of 2X Lysis Buffer (10% SDS, 100 mM TEAB pH 7.5, 2 mM MgCl2, and 2X HALT protease and phosphatase inhibitors). To obtain the 9, 30, and 300 sec time points, 80 µL of parasites in 1.5 mL tubes were incubated in a ThermoMixer (Eppendorf) set to 37°C. The parasites were stimulated by adding 20 µL of warmed 5X zaprinast (500 µM zaprinast in FluoroBrite) or a vehicle of 5X DMSO. At each time point, 20 µL of stimulated parasites were transferred directly into 20 µL of 2X Lysis Buffer to quench the reaction. Complete time courses were collected sequentially in the following order: auxin-treated stimulated with zaprinast, vehicle-treated stimulated with zaprinast, auxin-treated stimulated with DMSO, and vehicle-treated stimulated with DMSO. Lysates were treated with benzonase at a final concentration of 5 units/µL to remove DNA and were immediately subjected to protein cleanup and digestion. Protein cleanup and digestion Proteins were prepared for mass spectrometry using a modified version of the S-trap protocol (Protifi). Proteins in lysates were reduced with 5 mM TCEP for 10 min at 55°C and alkylated with 15 mM MMTS for 10 min at room temperature. The lysates were acidified to a final concentration of 1.2% v/v phosphoric acid. A 6X volume of S-trap binding buffer (90% methanol, 100 mM TEAB, pH 7.55) was mixed to each sample to precipitate proteins. The solution was loaded onto S-trap micro columns (Protifi) and spun at 4,000 x g for 1 min until all the solution had passed through the column. The columns were washed four times with 150 µL of S-trap binding buffer and centrifuged at 4,000 x g for 1 min between each wash. Proteins were digested on-column with 0.75 µg of trypsin (Promega) in 50 mM TEAB pH 8.5 overnight at 37°C in a humidified incubator. Digested peptides were eluted in three steps at 4,000 x g for 1 min: 40 µL of 50 mM TEAB, 40 µL of 0.2% formic acid, and 35 µL of 50% acetonitrile/0.2% formic acid. The peptide concentrations of eluted peptides were quantified using the Pierce Fluorometric Peptide Assay (Thermo Fisher Scientific) according to manufacturer’s instructions. The remaining samples were frozen in liquid nitrogen and lyophilized. TMTpro labeling Lyophilized peptides were resuspended in 100 mM TEAB pH 8.5 to peptide concentrations of 1.6 µg/µL. TMTpro reagents (Thermo Fisher Scientific; A44522 LOT# VI306829) were resuspended in acetonitrile to 25 µg/µL. 80 µg of peptides in 50 µL were combined with 250 µg of TMTpro reagent in 10 µL to achieve approximately a 3:1 label:peptide weight/weight ratio (Zecha et al. 2019). The TMTpro labels were assigned to minimize reporter ion interference and inter batch variability in the following scheme: Replicate 1 auxin: 0, 9, 30, 300 sec (126, 128C 130C, 132C); Replicate 1 vehicle: 0, 9, 30, 300 sec (127N, 129N, 131N, 133N); Replicate 2 auxin: 0, 9, 30, 300 sec (127C, 129C, 131C, 133C); Replicate 2 vehicle: 0, 9, 30, 300 sec (128N, 130N, 132N, 134) (Brenes et 56 al. 2019). The zaprinast and DMSO samples were incubated for 1 hr at room temperature shaking at 400 rpm. Unreacted TMTpro reagent was quenched with hydroxylamine at a final concentration of 0.2%. The samples were pooled, acidified to 3% with formic acid, and were processed using the EasyPep Maxi Sample Prep column (Thermo Fisher Scientific) according to the manufacturer’s instructions. 5% of the eluate volume was reserved as the unenriched proteome sample. The remaining eluted peptides were frozen in liquid nitrogen and lyophilized. Phosphopeptide enrichment Phosphopeptides were enriched using the Sequential enrichment from Metal Oxide Affinity Chromatography (SMOAC) protocol according to manufacturer instructions (Tsai et al. 2014). First, the High-Select TiO2 Phosphopeptide Enrichment Kit (Thermo Fisher Scientific) was used to enrich phosphopeptides from lyophilized TMTpro-labeled samples. The flow-through and contents of the first wash were pooled, frozen in liquid nitrogen, and lyophilized, along with the eluate. Second, the High-Select Fe-NTA Phosphopeptide Enrichment Kit (Thermo Fisher Scientific) was used to enrich phosphopeptides from the pooled flow-through and first wash from the previous enrichment. The eluted phosphopeptides were frozen in liquid nitrogen and lyophilized. Fractionation The enriched and unenriched samples were fractionated with the Pierce High pH Reversed-Phase Peptide Fractionation Kit (Thermo Fisher Scientific) according to manufacturer instructions for TMT-labeled peptides. The acetonitrile wash was omitted for enriched samples to prevent loss of phosphopeptides. The eluted peptides from the High-Select TiO2 Phosphopeptide Enrichment and High-Select Fe-NTA Phosphopeptide Enrichment were pooled prior to fractionation. 100 µg of unenriched samples were fractionated. 8 fractions were collected for each TMTpro set: zaprinast phosphoproteome (enriched) [1], zaprinast proteome (unenriched) [2], DMSO phosphoproteome (enriched) [3], and DMSO proteome (unenriched) [4]. The fractions were frozen in liquid nitrogen and lyophilized. MS data acquisition Lyophilized peptides were resuspended in approximately 15 µL (enriched) or 50 µL (unenriched) of 0.1% formic acid and were analyzed on an Exploris 480 Orbitrap mass spectrometer equipped with a FAIMS Pro source (Bekker-Jensen et al. 2020) connected to an EASY-nLC 1200 chromatography system using 0.1% formic acid as Buffer A and 80% acetonitrile/0.1% formic acid as Buffer B. Peptides were loaded onto a heated analytical column (ES900, Thermo, PepMap RSLC C18 3 µm, 100 Å, 75 µm x 15 cm, 40°C) via trap column (164946, Thermo, Acclaim PepMap C18 3 µm, 100 Å, 75 µm x 20 mm nanoViper). Peptides were separated at 300 nL/min. Enriched samples were separated on a gradient of 5–20% B for 110 min, 20–28% B for 10 min, 28–95% B for 10 min, 95% B for 10 min, 95–2% B for 2 min, 2% B for 2 min, 2–98% B for 2 min, 98% B for 2 min, 98–2% B for 2 min, and 2% B for 2 min. Unenriched samples were separated on a gradient of 5–25% B for 110 min, 25–40% B for 10 min, 40–95% B for 10 min, 95% B for 10 min, 95–2% B for 2 min, 2% B for 2 min, 2–98% B for 2 min, 98% B for 2 min, 57 98–2% B for 2 min, and 2% B for 2 min. The orbitrap and FAIMS were operated in positive ion mode with a positive ion voltage of 1800V; with an ion transfer tube temperature of 270°C; using a standard FAIMS resolution and compensation voltage of -50 and -65V, an inner electrode temperature of 100°C and outer electrode temperature 80°C with 4.5 mL/min carrier gas. DDA analysis was performed with a cycle time of 1.5 sec. Full scan spectra were acquired in profile mode at a resolution of 60,000, with a scan range of 400–1400 m/z, 300% AGC target, maximum injection time of 50 msec, intensity threshold of 5 x 104, 2–5 charge state, dynamic exclusion of 30 sec, mass tolerance of 10 ppm, purity threshold of 70%, and purity window of 0.7. MS2 spectra were generated with a HCD collision energy of 32 at a resolution of 45,000 using TurboTMT settings with a first mass at 110 m/z, an isolation window of 0.7 m/z, 200% AGC target, and maximum injection time of 120 msec. Phosphoproteomic time course analysis Raw files were analyzed in Proteome Discoverer 2.4 (Thermo Fisher Scientific) to generate peak lists and protein and peptide IDs using Sequest HT (Thermo Fisher Scientific) and the ToxoDB release 49 GT1 protein database. The maximum missed cleavage sites for trypsin was limited to 2. Precursor and fragment mass tolerances were 10 ppm and 0.02 Da, respectively. The following modifications were included in the search: dynamic oxidation (+15.995 Da; M), dynamic phosphorylation (+79.966 Da; S,T,Y), dynamic acetylation (+42.011 Da; N-terminus), static TMTpro (+304.207 Da; any N-terminus), static TMTpro (+304.207 Da; K), and static methylthio (+45.988 Da; C). TMTpro 16plex isotope correction values were accounted for (Thermo Fisher Scientific; A44522 LOT# VI306829). Peptides identified in each sample were filtered by Percolator to achieve a maximum FDR of 0.01 (Käll et al. 2007, 2008; Käll, Storey, and Noble 2008). Site localization scores were generated using ptmRS, with phosphoRS and use of diagnostic ions set to true (Taus et al. 2011). Reporter ion quantification used an integration tolerance of 20 ppm on the most confident centroid. For reporter ion quantification, unique peptides were quantified using a co-isolation threshold of 50, and average reporter signal-to-noise ratio of 10. Abundances were normalized on the total protein amount. Protein level and peptide level ratios were generated for each time point relative to 0 sec vehicle-treated parasites stimulated with DMSO. Exported peptide and protein abundance files from Proteome Discoverer 2.4 were loaded into R (version 4.1.1). To determine CDPK1-dependent phosphorylation, log2 ratios for peptide abundances in DMSO and zaprinast-treated samples derived from Proteome Discoverer were used. Only phosphorylated peptides quantified across all time points were used for analysis. Area under the curve (AUC) values were calculated for individual peptides undergoing vehicle (AUCveh) and auxin (AUCauxin) treatment using trapezoidal integration. AUC differences (AUCdiff) were calculated by taking the difference between AUCveh and AUCauxin values. The distribution of zaprinast AUCdiff values were tested against a null distribution derived from the DMSO AUCdiff values to calculate z- scores and p-values using a two-tailed t-test. Replicates 1 and 2 were analyzed independently and phosphopeptides with p-values < 0.05 across both replicates were determined to be CDPK1-dependent (Group A). Peptides exhibiting phosphorylation 58 independent of CDPK1 (Group B, C, and D) were determined similarly, but compared the distribution of zaprinast AUCvehicle values to a null distribution of DMSO AUCvehicle values and excluded phosphopeptides already determined to be CDPK1-dependent. CDPK1-independent phosphopeptides were clustered into Groups B, C, and D using projection-based clustering (Thrun and Ultsch 2021). Gene ontology enrichment Gene ontology terms were obtained for all genes present in the enriched zaprinast phosphoproteome from ToxoDB.org (Computed GO function and GO function IDs). Gene ontology IDs within each group of zaprinast-dependent genes (Groups A-D) were tested for enrichment against the entire enriched zaprinast phosphoproteome. Enrichment p values were generated using a hypergeometric test. Enrichment ratios were calculated by dividing the gene ratio (overlap/signatures) by the relative frequency of gene sets (gene sets/background). Only GO IDs with significant enrichment and an overlap of 2 were plotted. Thiophosphorylation enrichment was performed similarly. Metabolic tagging, click reaction, pull down, and Western blotting Metabolic tagging and cell lysis Upon infection of HFF monolayers the medium was removed and replaced by fresh culture media supplemented with 25 μM YnMyr (Iris Biotech) or Myr (Tokyo Chemical Industry). The parasites were then incubated for 16 hrs, washed with PBS (2x) and lysed on ice using a lysis buffer (PBS, 0.1% SDS, 1% Triton X-100, EDTA-free complete protease inhibitor (Roche Diagnostics)). Lysates were kept on ice for 20 min and centrifuged at 17,000 × g for 20 min to remove insoluble material. Supernatants were collected and protein concentration was determined using a BCA protein assay kit (Pierce). Click reaction and pull down Lysates were thawed on ice. Proteins (100–300 μg) were taken and diluted to 1 mg/mL using the lysis buffer. A click mixture was prepared by adding reagents in the following order and by vortexing between the addition of each reagent: a capture reagent (stock solution 10 mM in water, final concentration 0.1 mM), CuSO4 (stock solution 50 mM in water, final concentration 1 mM), TCEP (stock solution 50 mM in water, final concentration 1 mM), TBTA (stock solution 10 mM in DMSO, final concentration 0.1 mM) (Heal et al. 2011). Capture reagent used herein was the Trypsin cleavable reagents (Broncel et al. 2020). Following the addition of the click mixture the samples were vortexed (room temperature, 1 hr), and the reaction was stopped by addition of EDTA (final concentration 10 mM). Subsequently, proteins were precipitated (chloroform/methanol, 0.25:1, relative to the sample volume), the precipitates isolated by centrifugation (17,000 x g, 10 min), washed with methanol (1 x 400 μL) and air dried (10 min). The pellets were then resuspended (final concentration 1 mg/mL, PBS, 0.4 % SDS) and the precipitation step was repeated to remove excess of the capture reagent. Next, samples were added to 15 μL of pre-washed (0.2 % SDS in PBS (3 x 500 μL)) Dynabeads® MyOneTM Streptavidin C1 (Invitrogen) and gently vortexed for 90 min. The 59 supernatant was removed, and the beads were washed with 0.2 % SDS in PBS (3 x 500 μL). SDS-PAGE and Western blotting Beads were supplemented with 2% SDS in PBS (20 μL) and 4x SLB (Invitrogen), boiled (95°C, 10 min), centrifuged (1,000 x g, 2 min) and loaded on 10% or 4–20% SDS-PAGE gel (Bio-Rad). Following electrophoresis (60 min, 160V), gels were briefly washed with water and proteins were transferred (25 V, 1.3 A, 7 min) onto nitrocellulose membranes (Bio-Rad) using Bio-Rad Trans Blot Turbo Transfer system. After brief wash with PBS-T (PBS, 0.1% Tween-20) membranes were blocked (5% milk, TBS-T, 1h) and incubated with primary antibodies (5% milk, TBS-T, overnight, 4°C) at the following dilutions: rat anti-HA (1:1000; Roche Diagnostics), mouse anti-Myc (1:1000; Millipore), rabbit anti- Gra29 (1:1000; Moritz Treeck Lab), rabbit anti-SFP1 (1:1000; Moritz Treeck Lab), mouse anti-T. gondii [TP3] (1:1000; Abcam), mouse anti-CDPK1 (1:3000; John Boothroyd Lab), rabbit anti-SAG1 (1:10,000; John Boothroyd Lab), rabbit anti-GAP45 (1:1000; Peter Bradley Lab), mouse anti-GFP (1:1000, Roche Diagnostics) and rabbit anti-mCherry (1:1000, Abcam). Following washing (TBS-T, 3x) membranes were incubated with IR dye-conjugated secondary antibodies (1:10,000, 5% milk, TBS-T, 1 hr) and after a final washing step imaged on a LiCOR Odyssey imaging system (LI-COR Biosciences). MS detection of myristoylated CDPK1 Mass spectrometry proteomics methods and data for myristoylated CDPK1 are available from the ProteomeXchange Consortium via the PRIDE partner repository (ID PXD019677) and the associated publication (Broncel et al. 2020). Depletion of mAID tagged CDPK1 (iKD) Parasites were treated with 500 µM auxin or equivalent volume of vehicle (ethanol) for at least 2 hrs prior to Western blot analysis. Subcellular fractionation RH ∆ku80∆hxgprt YFP expressing parasites were metabolically tagged with 25 μM Myr or YnMyr for 16 hrs. Following a PBS wash, the parasites were syringe lysed in Endo buffer (44.7 mM K2SO4, 10 mM MgSO4, 106 mM sucrose, 5 mM glucose, 20 mM Tris– H2SO4, 3.5 mg/mL BSA, pH 8.2) and collected by centrifugation (512 x g, 10 min). The parasites were then lysed in 300 μL of cold hypotonic buffer (10 mM HEPES, pH 7.5) supplemented with protease inhibitors (Roche), passed through 25G needle (5x) and left on ice for 40 min. Next, lysates were pelleted by centrifugation (16,000 x g, 20 min, 4 °C) and the resulting supernatant was subjected to an additional high speed (100,000 x g, 1 h, 4 °C) centrifugation step. To avoid the loss of the high-speed pellet, only half of the supernatant was removed at this point. Each fraction was then taken up in 0.4% (final) SDS HEPES, clicked to a capture reagent and pulled down as described above. Myristoylation-dependent partitioning was revealed by SDS-PAGE and Western blotting. 60 Myristoylation-dependent fractionation for CDPK1 complemented WT and Mut lines: parasites were seeded 24 hrs prior experiment. Following a PBS wash, the parasites were syringe lysed in Endo buffer (44.7 mM K2SO4, 10 mM MgSO4, 106 mM sucrose, 5 mM glucose, 20 mM Tris–H2SO4, 3.5 mg/mL BSA, pH 8.2) and collected by centrifugation (512 x g, 10 min). The parasites were then lysed in 300 μL of cold hypotonic buffer (10 mM HEPES, pH 7.5) supplemented with protease inhibitors (Roche), passed through 25G needle (5x) and left on ice for 40 min. Next, lysates were pelleted by centrifugation (100,000 x g, 1 hr, 4 °C), the supernatant was removed, and cytosolic proteins precipitated with methanol/chloroform. Proteins from the pellet and supernatant fractions were dissolved in 2% (final) SDS PBS and myristoylation-dependent partitioning was revealed by SDS-PAGE and Western blotting. Immunoprecipitation of cWT and cMUT CDPK1 Parasite harvest cWT, cMUT, and TIR1 (untagged) parasites were infected onto confluent HFFs in 15 cm dishes. At 1-day post-infection (dpi), 50 µM auxin or vehicle was added to deplete endogenous mAID-tagged CDPK1 and 1 µM compound 1 was added to block egress until parasites were ready to harvest. At 2 dpi, infected HFFs were washed twice with PBS to wash out drugs. Parasites were mechanically released in Endo buffer with a 27G needle, passed through a 5 µm filter, and spun at 1000 x g for 10 min. Parasite pellets were resuspended in a cold hypotonic buffer with 1X HALT protease inhibitors to parasite concentrations of 1.1 x 109 tg/mL, passed through a 27G needle five times, and incubated on ice for 1 hr to complete hypotonic lysis. The samples were spun at 1000 x g for 5 min to pellet unlysed parasites and the supernatant was saved. NaCl was added to a final concentration of 150 mM and this was used as the immunoprecipitation input. Immunoprecipitation 25 µL of anti-HA magnetic beads (Thermo) were used per condition. Beads were washed twice with a wash buffer (10 mM HEPES, 150 mM NaCl, pH 7.5). To begin pulldown, parasite lysate was used to resuspend washed beads and incubated for 1 hr rotating at room temperature. Beads were washed four times with a wash buffer. Proteins were eluted by resuspending beads in 20 µL of 1X S-trap sample buffer (5% SDS, 50 mM TEAB, pH 7.5) and incubated at 70°C for 10 min. The eluate was collected for MS sample processing and analysis. Results are representative of two independent experiments. Protein cleanup and digestion Proteins were prepared for mass spectrometry as described above in “Sub-minute phosphoproteomics - Protein cleanup and digestion”. Eluted peptides were frozen in liquid nitrogen, lyophilized, and stored at -80°C until MS analysis. MS data acquisition Lyophilized peptides were resuspended in 20 µL of 0.1% formic acid and were analyzed on an Exploris 480 Orbitrap mass spectrometer equipped with a FAIMS Pro source (Bekker-Jensen et al. 2020) connected to an EASY-nLC chromatography system using 61 0.1% formic acid as Buffer A and 80% acetonitrile/0.1% formic acid as Buffer B. Peptides were separated at 300 nL/min on a gradient of 1–6% B for 1 min, 6–21% B for 41 min and 30 sec, 21–36% B for 20 min and 45 sec, 36–50% B for 10 min and 15 sec, 100% B for 14 min and 30 sec, 100–2% B for 3 min, 2% B for 3 min, 2–98% B for 3 min, and 98% B for 3 min. The orbitrap and FAIMS were operated in positive ion mode with a positive ion voltage of 1800V; with an ion transfer tube temperature of 270°C; using a standard FAIMS resolution and an inner and outer electrode temperature of 100°C with 4.5 mL/min carrier gas. Samples were analyzed twice in DIA mode with a compensation mode of -50 and -65V. Full scan spectra were acquired in profile mode at a resolution of 120,000, with a scan range of 400–1000 m/z, 300% AGC target, and auto mode for maximum injection time. MS2 spectra for the DIA scan were generated with a isolation window of 20 m/z with a 0 m/z window overlap, 30 scan events, a HCD collision energy of 30 at a resolution of 30,000, first mass at 200 m/z, precursor mass range of 400–1000 m/z, and a standard AGC target and automatically determined maximum injection time. Immunoprecipitation data analysis DIA-MS samples were analyzed using Scaffold DIA (2.0.0). DIA-MS data files were converted to mzML format using ProteoWizard (3.0.19254) (Chambers et al. 2012). Analytic samples were aligned based on retention times and individually searched against dku80_FAIMS_DIA_90min_autoIT.blib with a peptide mass tolerance of 10 ppm and a fragment mass tolerance of 0.02 Da. The ToxoDB release 46 GT1 protein database was used for protein identification. The following modifications were included in the search: dynamic oxidation (+15.995 Da; M), dynamic phosphorylation (+79.966 Da; S,T,Y), and static methylthio (+45.988 Da; C). The digestion enzyme was trypsin with a maximum of 2 missed cleavage sites allowed. Only peptides with charges in the range of 2 to 3 and length in the range 6 to 30 were considered. Peptides identified in each sample were filtered by Percolator to achieve a maximum FDR of 0.01 (Käll et al. 2007, 2008; Käll, Storey, and Noble 2008). Individual search results were combined, and peptide identifications were assigned posterior error probabilities and filtered to an FDR threshold of 0.01 by Percolator. Peptide quantification was performed by Encyclopedia (0.9.2). For each peptide, the 5 highest quality fragment ions were selected for quantification. Proteins that contained similar peptides and could not be differentiated based on MS/MS analysis were grouped to satisfy the principles of parsimony. Protein groups were thresholded to achieve a protein FDR less than 1%. Significance values were derived from t-tests across two replicates and adjusted with Benjamini-Hochberg correction with an FDR of 0.05. Exported protein abundance files from Scaffold DIA were loaded into R (version 4.1.1). Thiophosphorylation of CDPK1 substrates Parasite harvest and treatment T. gondii tachyzoites from the RH strain (CDPK1G and CDPK1M) were passaged twice across 4 days in SILAC media in T12.5 flasks. CDPK1G parasites were grown in “heavy” SILAC media (DMEM 88364, 10% dialyzed FBS, 0.1 mg/mL 13C15N L-arginine, 0.1 mg/mL 13C15N L-lysine) and CDPK1M parasites were grown in “light” SILAC media 62 (DMEM 88364, 10% dialyzed FBS, 0.1 mg/mL L-Arginine, 0.1 mg/mL L-Lysine). On day 4, confluent HFFs in 15 cm dishes were infected with CDPK1G and CDPK1M parasites in SILAC media. On day 6, extracellular parasites were harvested by filtering through a 5 µm filter and pelleted at 1000 x g for 7 min at 4°C. Parasites were washed once in 1X intracellular buffer (ICB) (137 mM KCl, 5 mM NaCl, 20 mM HEPES, 10 mM MgCl2, pH 7.5 KOH) and then resuspended in 400 µL of 1X ICB. Parasites were semi-permeabilized after the addition of 400 µL of 6 µg/mL aerolysin diluted in 1X ICB and incubated at 37°C for 10 min. After semi-permeabilization, 400 µL of 4X Ca2+ solution (16 mM CaEGTA, 100 ng/mL 1B7 nanobody in 1X ICB) followed by 400 µL of 4X ATP solution (4 mM GTP, 0.4 mM ATP, 0.2 mM KTPγS, 1X HALT protease and phosphatase inhibitor in 1X ICB). The kinase reaction was initiated by incubating parasites at 30°C for 5 min. Parasites were pelleted at 1000 x g at 4°C for 10 min. Parasite pellets were resuspended in 250 µL of 1X lysis buffer (10% TritonX-100, 1X HALT protease and phosphatase inhibitor, 10 mM K2EGTA in 1X ICB). Protein quantification and thiophosphorylation immunoblotting Proteins in lysates were quantified using DC assay (BioRad) utilizing BSA as a protein standard and a diluent (150 mM NaCl, 20 mM Tris pH 7.6) to prepare standard curves and dilution series. Thiophosphorylation was verified using immunoblot by first incubating 3 µL of sample with p-nitrobenzyl mesylate diluted in 1X ICB at a final concentration at 2 mM for 2 hrs at room temperature. 5X Laemmli sample buffer was added (see “Immunoblotting” for recipe) and samples were boiled for 10 min. Samples were resolved on a home-made polyacrylamide gel (5% stacking, 15% resolving, 15- well, 0.75 mm) and transferred overnight at 4°C. Nitrocellulose membranes were blocked with 5% milk in TBS-T for 1 hr. Primary antibody incubations were performed with an anti-thiophosphate ester antibody (rabbit 51-8; 1:5000) and anti-tubulin (mouse 12G10; 1:2000) for 1 hr at room temperature. Secondary incubations were performed with LI- COR antibodies (rabbit 680, mouse 800; 1:10,000) for 1 hr at room temperature. Imaging was performed using a LI-COR Odyssey. Protein cleanup and digestion Proteins were precipitated using methanol chloroform extraction. For 250 µL of lysate, 800 µL methanol and 200 µL of chloroform was added, followed by vortexing. After adding 600 µL of water, the sample was vortexed and centrifuged at max speed for 5 min at 4°C. After removing the supernatant without disrupting the precipitate, 375 µL of methanol was added and vortexed. Samples were centrifuged at max speed for 15 min at 4°C. The protein pellet was allowed to air dry after removing the supernatant. Dried protein pellets were resuspended in 200 µL of 8M urea. 1 mg of protein (determined from protein quantification; see above) from CDPK1G and CDPK1M parasites were pooled for a total of 2 mg of protein. To digest protein, 5X volume of 1X Trypsin Digest Buffer (100 mM ammonium acetate pH 8.9, 1mM CaCl2, 2 mM TCEP) was added to the sample followed by 40 µg of sequencing grade trypsin (Promega). Proteins were digested overnight rotating at room temperature. To prepare samples for desalting, glacial acetic acid was added to 10% (v/v) and debris was briefly spun down. A C18 Sep-Pak Plus cartridge (Waters) was prepared using a syringe pump and 10 mL syringe with three 10 63 mL washes at a flow rate of 2 mL/min: 0.1% acetic acid, 90% acetonitrile/0.1% acetic acid, and 0.1% acetic acid. The sample was loaded in a 5 mL syringe at a flow rate of 0.5 mL/min. The syringe and cartridge were washed with 5 mL of 0.1% acetic acid at a flow rate of 0.5 mL/min. A final wash was performed using 10 mL of 0.1% acetic acid at a flow rate of 2 mL/min. Peptides were eluted with 4.5mL of 40% acetonitrile/0.1% acetic acid at a flow rate of 0.5 mL/min. Samples were spun in a speed vac for 3 hrs until samples could be pooled into a single 2 mL tube. Samples were frozen in liquid nitrogen and lyophilized overnight. Lyophilized peptides were stored at -80°C. Thiophosphate enrichment To enrich for thiophosphorylated peptides, 400 µL of SulfoLink Coupling Resin slurry was used (Thermo). Incubations were performed in the dark using aluminum foil due to the light sensitivity of the resin. The resin was washed twice with 1 mL of binding buffer (25 mM HEPES, 50% acetonitrile, pH 7.0 NaOH + HCl) rotating for 5 min followed by a 1000 rpm spin for 10 sec. The supernatant was removed without disturbing the pelleted resin. The beads were blocked with 1mL of blocking buffer (binding buffer with 25 µg/mL β-casein and 2 mM TCEP) and rotated for 5 min at room temperature. After pelleting the resin and removing the supernatant, the resin was resuspended with lyophilized peptides dissolved in 500 µL of the blocking buffer. Samples were incubated overnight at room temperature. Samples were spun at 1000 rpm for 10 sec and the supernatant was spun again, lyophilized, and stored at -80°C prior to MS analysis. The beads received the following series of 1 mL washes for 5 min rotating and spun at 1000 rpm for 10 sec: twice with binding buffer with 2 mM TCEP, once with quenching buffer (25 mM HEPES, 50% acetonitrile, 5 mM DTT, pH 8.5 NaOH), once with binding buffer with 2 mM TCEP, once with 5% formic acid (no rotation), once with binding buffer with 2 mM TCEP, and three times with 0.1% acetic acid. The resin was no longer light sensitive after incubation with the quenching buffer. To elute captured peptides, resin was resuspended in 500 µL OXONE (2 mg/mL potassium monopersulfate) and rotated for 5 min at room temperature. The supernatant containing eluted peptides was collected after spinning at 1000 rpm for 10 sec. OXONE was removed from the sample using C18 spin columns (Pierce) according to manufacturer instructions. A total of 4 washes with the equilibrium/wash buffer was performed. Peptides were spun in a speed vac until dry and stored at -80°C until MS analysis. MS data acquisition The samples were resuspended in 10–20 µL of 0.1% formic acid for MS analysis and were analyzed on a Q-Exactive HF-X Orbitrap mass spectrometer connected to an EASY-nLC 1200 chromatography system using 0.1% formic acid as Buffer A and 80% acetonitrile/0.1% formic acid as Buffer B. Peptides were loaded onto a analytical column (column: PF360-75-15-N-5, New Objective, 360 µm OD, 75 µm ID, 15 µm Tip PicoFrit Emitter; resin: 04A-4506, Phenomenex Aeris Peptide, C18 1.7 µm) via trapping column (column: 360 um OD, 100 um ID; resin: AA12S11, YMC Gel ODS-A, C18 10 µm). Peptides were separated at 300 nL/min on a gradient of 2% B for 5 min, 2–25% B for 100 min, 25–40% B for 20 min, 40–100% B for 1 min, and 100% B for 12 min. The orbitrap was operated in positive ion mode with a positive ion voltage of 2700V with an ion transfer 64 tube temperature of 300°C. Full scan spectra were acquired in profile mode at a resolution of 60,000, with a scan range of 375 to 1600 m/z, 1 x 106 AGC target, maximum injection time of 50 msec, intensity threshold of 4 x 105, dynamic exclusion of 13 sec, and 20 data dependent scans (DDA Top 20). MS2 spectra were generated with a HCD collision energy of 27 at a resolution of 15,000, first mass at 100 m/z, isolation window of 1.5 m/z, an AGC target of 1 x 105 with a maximum injection time of 20 msec, and scan range of 200 to 2000 m/z. Thiophosphorylation analysis Raw files were analyzed in Proteome Discoverer 2.2 (Thermo Fisher Scientific) to generate peak lists and protein and peptides identifications using Sequest HT (Thermo Fisher Scientific) and the ToxoDB release 34 GT1 protein database. The maximum missed cleavage sites for trypsin was limited to 2. The following modifications were included in the search: dynamic oxidation (+15.995 Da; M), dynamic phosphorylation (+79.966 Da; S,T,Y), dynamic 13C 15N (+10.008 Da; R), dynamic 13C 156 4 6 N2 (+8.014 Da; K), and dynamic acetylation (+42.011 Da; N-terminus). Site localization scores were generated using ptmRS, with phosphoRS and use of diagnostic ions set to true. SILAC 2plex (Arg10, Lys8) method was used for relative quantification of protein and unique peptide abundances. For peptide level analysis, ratios of unique peptides comparing CDPK1G and CDPK1M were generated and low abundance resampling (5%) was used to impute missing values. For protein level analysis, ratios of proteins were determined from the summed abundance of unique peptides comparing CDPK1G and CDPK1M strains. Peptide and protein level data were exported from Proteome Discoverer for analysis in R. For enriched peptides, abundance ratios were calculated within each replicate for high confidence peptides (CDPK1G/CDPK1M) and normalized to the median abundance ratio of the whole proteome peptides derived from the flow through samples. Normalized abundance ratios in the flow through proteome samples were calculated by dividing abundance ratios by the median abundance ratios within each replicate. For enriched peptides, an average log2 normalized abundance ratio was calculated from three replicates. Significantly enriched peptides were calculated using a one-tailed t-test using the normalized abundance ratios for three replicates. A non-linear significance threshold was calculated using the function (y = |4/x|), where significantly enriched peptides had product value of -log10(p)*(mean log2 abundance ratio) greater than 4. Immunoblotting Samples were combined with 5X Laemmli sample buffer (10% SDS, 50% glycerol, 300mM Tris HCl pH 6.8, 0.05% bromophenol blue, 5% beta-mercaptoethanol) and were incubated at 95°C for 10 min. The samples were run on precast 4–15% or 7.5% SDS gels (Bio-Rad) and were transferred overnight onto a nitrocellulose membrane in transfer buffer (25mM TrisHCl, 192 mM glycine, 0.1% SDS, 20% methanol) at 4°C. Blocking was performed with 5% milk in PBS for 1 hr rocking at room temperature. Antibody incubations were performed with 5% milk in TBS-T for 1 hr rocking at room temperature. Three 5 min TBS-T washes were performed before and after secondary antibody 65 incubations rocking at room temperature. After a final PBS wash, imaging was performed using a LI-COR Odyssey. For immunoblot detection of the HA tag in AID-HOOK and FTS-AID after auxin-mediated depletion, secondary antibody incubations were performed with anti-rabbit HRP antibodies (Jackson ImmunoResearch) and detected with chemiluminescence (Azure) for increased sensitivity. Imaging was performed using a Bio-Rad Gel Doc XR. Immunofluorescence analysis Parasite-infected HFF monolayers grown on glass coverslips were fixed with 3% formaldehyde for 15 min prior to washing with PBS. Fixed cells were then permeabilized (PBS, 0.1% Triton X-100, 10 min), blocked (3% BSA in PBS, 1 hr), and labeled with anti- HA (1:1000, 1hr; Roche). HA-tagged CDPK1 in the cWT and cMut lines was visualized with secondary goat antibodies (1:2000, 1 hr; Life Technologies) conjugated to Alexa Fluor 594 and 488, respectively. Cytosolic GFP (cWT) and mCherry (cMut) were used as parasite counterstains. Nuclei were visualized with the DNA stain (DAPI; Sigma) added at 5 µg/mL with the secondary antibody. Stained coverslips were mounted on glass slides with Slowfade (Life Technologies) and imaged on a Nikon Eclipse Ti-U inverted fluorescent microscope. Images were analyzed using Nikon NIS Elements imaging software. Plaque Assays Parasites were harvested by syringe lysis, counted, and 200 parasites were seeded on confluent HFF monolayers grown in 24-well plates (Falcon). Wells were treated with 500 μM auxin or vehicle (ethanol) and plaques were allowed to form for 5 days. Plaque formation was assessed by inspecting the methanol fixed and 0.1% crystal violet stained HFF monolayers. Egress assays Parasites were added to HFF monolayer and grown for 24 hrs in a 96 well plate. The wells were then treated with 500 μM auxin or an equivalent volume of vehicle (ethanol) for 2 hrs and then washed with PBS (2x). The media was exchanged for 100 μl Ringer’s solution (155 mM NaCl, 3 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 3 mM NaH2PO4, 10 mM HEPES, 10 mM glucose) and the plate was placed on a heating block to maintain the temperature at 37°C. To artificially induce egress, 50 μL of Ringer’s solution containing 24 μM ionophore (8 μM final, Thermo) was added to each well. At specified time points the cells were fixed by adding 33 μL 16% formaldehyde (3% final) for 15 min. Cells were washed in PBS (3x) and stained with rabbit anti-TgCAP 1:2000 (Hunt et al. 2019) followed by goat anti-rabbit Alexa Fluor 488 (1:2000) and DAPI (5 μg/mL). Automated image acquisition of 25 fields per well was performed on a Cellomics Array Scan VTI HCS reader (Thermo Scientific) using a 20x objective. Image analysis was performed using the Compartmental Analysis BioApplication on HCS Studio (Thermo Scientific). Egress levels were determined in triplicate for three independent assays. Vacuole counts were 66 normalized to t = 0 to determine how many intact vacuoles had remained after egress. The results were statistically tested using one-way ANOVA with Tukey’s multiple comparison test in GraphPad Prism 7. The data are presented as mean ± s.d. AUTHOR CONTRIBUTIONS Alex W Chan Contribution: Conceptualization, Methodology, Validation, Formal analysis, Investigation, Writing - original draft, Writing - review and editing, Visualization Malgorzata Broncel Contribution: Methodology, Validation, Formal analysis, Investigation Nicole Haseley Contribution: Methodology, Formal analysis Alice L Herneisen Contribution: Resources, Methodology Emily Shortt Contribution: Resources Moritz Treeck Contribution: Supervision, Resources, Funding acquisition, Writing - review and editing Sebastian Lourido Contribution: Conceptualization, Resources, Supervision, Funding acquisition, Methodology, Writing - review and editing 67 CHAPTER 3: The HOOK complex promotes microneme exocytosis to enable invasion Alex W Chan1,2, Nicole Haseley1, Sundeep Chakladar1, Elena Andree1, Alice L Herneisen1,2, Sebastian Lourido1,2 1 Whitehead Institute for Biomedical Research, Cambridge, MA, USA 2 Biology Department, Massachusetts Institute of Technology, Cambridge, MA, USA The following chapter is adapted from an article published in eLife (Chan et al., 2023). INTRODUCTION Combining sub-minute resolution phosphoproteomics and bio-orthogonal labeling of direct kinase substrates enabled the identification of CDPK1 substrates that may regulate microneme exocytosis, but these findings required the identification and characterization and a new regulator of microneme exocytosis. To identify new factors involved in Ca2+-regulated secretion, we prioritized Class 1 and Class 2 candidates. Seven candidates in this category lack functional annotation or have been associated with apical structures: TGGT1_227610, TGGT1_221470, TGGT1_235160, TGGT1_254870, KinesinB (TGGT1_273560), a small nuclease (TGGT1_310060), and TGGT1_289100 (MIC18). TGGT1_221470 was previously identified as a CDPK1 substrate after a pull down from parasite lysates but remained functionally uncharacterized (Lourido, Jeschke, et al. 2013). KinesinB localizes to cortical microtubules, TGGT1_254870 localizes to the apical complex, and TGGT1_289100 co- localizes with micronemes, each of which are parasite structures relevant to exocytosis (Butler et al. 2014; Leung et al. 2017; Long, Anthony, et al. 2017). Of these candidates, only TGGT1_227610 and TGGT1_289100 appear to be required for parasite fitness (Sidik, Huet, et al. 2016b; Long, Anthony, et al. 2017). Despite prior annotation as a microneme protein, TGGT1_289100 lacks a signal peptide and is localized to the cytosol by spatial proteomics (Butler et al. 2014; Barylyuk et al. 2020). Instead, TGGT1_289100 is predicted to have an N-terminal Hook domain and extensive coiled-coil domains (Söding, Biegert, and Lupas 2005; Simm, Hatje, and Kollmar 2015) with homology to activating adaptors that bind to endosomes and activate super-processive dynein- 68 mediated trafficking towards the minus end of microtubules (Figure 7A) (Bielska et al. 2014; Zhang et al. 2014; Guo et al. 2016). The predicted polarity of cortical microtubules and their association with micronemes support a model where the vesicles are trafficked by dyneins towards the apical end of the parasite (Chen et al. 2015; Leung et al. 2017; Wang et al. 2021). We observed extensive CDPK1-dependent phosphorylation between the Hook and coiled-coil domains of TGGT1_289100 (S167, S177, and S189–191), consistent with potential regulation of the putative adaptor. These data motivated functional characterization of this factor—henceforth referred to as HOOK. RESULTS A Hook domain protein phosphorylated by CDPK1 regulates parasite invasion To study the role of HOOK during the acute stages of the parasite, we generated a conditional knockdown by fusing an HA-mAID tag to its N terminus (AID-HOOK; Figure 8A). HOOK was depleted from parasites following 24–40 hrs of auxin treatment, as determined by immunoblotting or immunofluorescence microscopy (Figure 7B–C). In contrast to previous observations that localized HOOK to micronemes, we observed only partial co-localization with micronemes with a majority of HOOK localized to the cytosol (Figure 7C). Plaque formation was impaired when AID-HOOK parasites were grown in the presence of auxin, consistent with the strong effect on parasite fitness reported from genome-wide knockout screens (Figure 7D)(Sidik, Huet, et al. 2016b). These results indicate that HOOK is required during the acute stages of T. gondii. Parasite replication was not affected following 24 hrs of auxin treatment (Figure 7E). Microneme and rhoptry biogenesis were also unaffected following 24 hrs of auxin treatment immunofluorescence analysis (Figure 7C, Figure 8B). These data suggest HOOK functions during stages associated with parasite motility, such as parasite invasion and egress. Parasites depleted of HOOK displayed reduced invasion efficiency, consistent with that of CDPK1-depleted parasites (Figure 7F). We noted that even in the absence of auxin treatment AID-HOOK parasites required longer incubations to reach comparable 69 levels of invasion to wildtype, suggesting that manipulation of the N terminus partially affects HOOK function. HOOK depletion only partially recapitulates the effects of CDPK1 loss. Parasites depleted of CDPK1 were unable to egress—as documented previously (Lourido et al. 2010; Lourido, Tang, and Sibley 2012)—whereas depletion of HOOK had no effect on parasite egress (Figure 7G). Finally, we also examined whether the stability of CDPK1 affected HOOK expression by tagging the C terminus of HOOK with a 3xHA tag in Figure 7. HOOK is required for host cell invasion, but dispensable for egress. (A) Schematic of T. gondii and H. sapiens HOOK protein domains. HOOK domain (blue), coiled-coil domain (yellow), sites phosphorylated by CDPK1 (red). (B) Immunoblot of HOOK conditional knockdown parasites (AID- HOOK) after auxin treatment for 40 hrs compared to untagged TIR1 parasites. CDPK1 was used as a loading control. (C) AID-HOOK is visualized in fixed intracellular parasites by immunofluorescence after auxin treatment for 24 hrs. Hoechst and MIC2 are used as counterstains. (D) Plaque assays of host cells infected with TIR1 or AID-HOOK parasites for 8 days in auxin. Host cells are stained with crystal violet. (E) Replication assays of host cells infected with TIR1 or AID- HOOK parasites in auxin for 24 hrs. Parasites per vacuole were quantified from immunofluorescence on fixed intracellular parasites. p > 0.9. Two-way ANOVA. (F) Invasion assays of untagged TIR1, CDPK1-AID, and AID- HOOK parasites treated auxin for 40 hrs. Medians are plotted for n = 3 biological replicates (different shades of gray); n.s., p > 0.05, Welch’s t-test. (G) Parasite egress stimulated with zaprinast following treatment with auxin for 24 hrs. Egress was monitored by live microscopy. Percent egress plotted for n = 3 biological replicates, n.s., p > 0.05, Welch’s t-test. (H) HOOK tagged with a C- terminal 3xHA in CDPK1 cKD parasites (CDPK1- AID) visualized in fixed intracellular parasites by immunofluorescence as in D. 70 CDPK1 cKD parasites (HOOK-3xHA). HOOK localization and abundance was unaffected by depleting parasites of CDPK1 for 24 hrs as determined by immunofluorescence (Figure 7H). Taken together, our results indicate that HOOK is required for invasion of host cells, but dispensable for egress. Figure 8. Extended analysis of HOOK knockdown. (A) PCR analysis confirming correct integration of the HA-mAID cassette at the N terminus of endogenous HOOK (TGGT1_289100) in the TIR1 line. (B) Rhoptries (ROP1) are visualized in fixed intracellular parasites by immunofluorescence after treatment with auxin for 24 hrs. Hoechst and GAP45 are used as counterstains. HOOK forms a complex that regulates parasite invasion Opisthokont HOOK proteins function in complexes to activate dynein-mediated trafficking of vesicular cargo along microtubules (Bielska et al. 2014; Xu et al. 2008; Yao, Wang, and Xiang 2014; Guo et al. 2016; Gillingham et al. 2014; Christensen et al. 2021). In D. melanogaster and mammals, HOOK proteins have been shown to form dimers and bind FTS and FHIP via a C-terminal region that interacts with vesicular cargo (Christensen et al. 2021; Krämer and Phistry 1996; Xu et al. 2008; Lee et al. 2018). To identify analogous binding partners of HOOK in T. gondii, we performed IP-MS on lysates from HOOK-3xHA parasites and an untagged control (Figure 9A). Three proteins showed comparable enrichment to HOOK-3xHA: TGGT1_264050, TGGT1_316650, and TGGT1_306920. Like FTS, TGGT1_264050 bears homology to E2 ubiquitin-conjugating 71 enzymes, but lacks the catalytic cysteine required for enzymatic activity at position 162. Because of these shared features and their homology, we designated TGGT1_264050 as FTS. To confirm that FTS forms a complex with HOOK, we investigated its localization and performed reciprocal IPs. We fused a 3xHA tag to the C terminus of FTS and observed a similar localization to HOOK by immunofluorescence: primarily cytosolic with partial overlap with micronemes (Figure 9B). To identify binding partners of FTS, we performed IP-MS on FTS-3xHA parasite lysates and compared protein enrichment to the HOOK-3xHA IP (Figure 9C). Four proteins were significantly enriched in the reciprocal IP experiments—HOOK, FTS, TGGT1_316650, and TGGT1_306920— confirming the four-member complex. 72 Figure 9. The HOOK complex is required for microneme exocytosis. (A) IP-MS of HOOK-3xHA or untagged parasites. Protein abundances determined by LC-MS/MS are shown for n = 3 biological replicates. Significantly enriched proteins (red) based on more than 3 unique peptides and p < 0.05; ANOVA and Benjamini-Hochberg corrected. (B) FTS-3xHA visualized in fixed intracellular parasites by immunofluorescence after treatment with auxin for 24 hrs. Hoechst and MIC2 are used as counterstains. (C) Reciprocal IP-MS of HOOK-3xHA and FTS-3xHA. FTS is tagged with a C-terminal 3xHA epitope at the endogenous locus (FTS-3xHA). IP enrichment is shown as the fold-change of protein abundances in tagged versus untagged strains determined by LC-MS/MS across n = 3 biological replicates. Significantly enriched proteins (red), for proteins with more than 3 unique peptides, pHOOK < 0.05, and pFTS < 0.05; ANOVA and Benjamini-Hochberg corrected. (D) Immunoblot of FTS cKD parasites. FTS is tagged with an C-terminal mAID-HA at its endogenous locus (FTS-AID) and treated with auxin for 40 hrs. ALD is used as a loading control. (E) Plaque assays of host cells infected with TIR1 or FTS-AID parasites for 8 days in auxin. Host cells are stained with crystal violet. (F) Micronemes are visualized in fixed intracellular FTS-AID and TIR1 parasites by immunofluorescence after treatment auxin for 24 hrs. Hoechst and GAP45 are used as counterstains. (G) Invasion assays of untagged TIR1, AID-HOOK, and FTS-AID parasites treated auxin for 40 hrs. Medians are plotted for n = 3 biological replicates (different shades of gray), n.s., p > 0.05, Welch’s t-test. (H) Parasite egress stimulated zaprinast following auxin treatment for 24 hrs. Egress was monitored by live microscopy. Percent egress plotted for n = 3 biological replicates, n.s., p > 0.05, Welch’s t-test. (I) Proximity labeling MS of FTS using TurboID (FTS-TurboID) compared to a cytosolic TurboID control (cytosolic mNeonGreen-TurboID). Protein abundances determined by LC-MS/MS are shown for n = 3 biological replicates. Significantly enriched proteins in FTS-TurboID are colored in red (red and blue), unique peptides > 3, ratio > 1, p < 0.05, ANOVA and Benjamini-Hochberg corrected. (J) Microneme protein secretion assays of parasites treated with auxin for 40 hrs. Extracellular parasites are stimulated with 1% ethanol (EtOH) and 3% IFS for 1.5 hrs. Percent MIC2 secreted is plotted for n = 3 biological replicates, n.s., p > 0.05, Welch’s t-test. 73 To determine whether FTS functions in the same pathway as HOOK, we generated a conditional knockdown by fusing an AID-HA tag to the C terminus of FTS (FTS-AID; Figure 10A). FTS was readily depleted from parasites following 40 hrs of auxin treatment (Figure 9D). FTS depletion completely blocked plaque formation (Figure 9E). Similar to HOOK, depletion of FTS had no observable effect on parasite replication or microneme biogenesis (Figure 10C, Figure 9F). During the motile stages of the parasite, FTS depletion resulted in a block in invasion efficiency but had no effect on egress, phenocopying the effects of HOOK depletion (Figure 9G–H). FTS appeared to tolerate tagging better than HOOK, since the FTS-AID parasites behaved normally in the absence of auxin. These data suggest that HOOK and FTS form a functional complex required for parasite invasion. We pursued enzyme-catalyzed proximity labeling to complement IP-MS studies and capture transient protein interactions (Branon et al. 2018). In this approach, cells express a protein of interest tagged with the promiscuous biotin ligase TurboID. Addition of a biotin substrate to live cells results in biotinylation within a few nanometers of the TurboID-tagged protein, in a matter of minutes. Biotinylated proteins can be enriched from cellular lysates using streptavidin-affinity purification and identified using MS. Proximity labeling was recently used to characterize the cargo diversity of distinct human FHF complexes in human cells (Christensen et al. 2021). To identify additional proteins that interact with the T. gondii HOOK complex, we performed proximity labeling after fusing TurboID to the C terminus of FTS, which was more amenable to tagging compared to HOOK (Figure 9I, Figure 10E). As a control, we used parasites expressing a cytosolic mNeonGreen-TurboID that would broadly label cytosolic proteins and identify non- specific interactions (Figure 10D, Figure 10F). Of the 14 proteins significantly enriched in FTS-TurboID parasites, members of the HOOK complex (HOOK, FTS, TGGT1_306920, and TGGT1_316650) were the top four most enriched proteins. In humans, proximity labeling studies suggest FTS and FHIP bind at the C terminus of the HOOK dimer to mediate FHIP binding to RAB5 endosomes (Christensen et al. 2021). Our FTS proximity labeling results suggest that TGGT1_306920 and TGGT1_316650 may also bind in this manner. Among the other significantly- 74 enriched proteins, PC and ACC1 encode carboxylases known to be covalently modified by biotin and likely represent non-specific enrichment. DHFR enrichment is likely due to its use as a selectable marker when generating the FTS-TurboID strain. TGGT1_294610 and TGGT1_280770 are a putative histone lysine methyltransferase and regulator of chromosome condensation (RCC1)-repeat containing protein, respectively, and have no characterized functions. Of the remaining enriched proteins that lack annotations, TGGT1_221180 was previously localized to micronemes by spatial proteomics and contains a transmembrane domain, suggesting a molecular link between the HOOK complex and micronemes (Barylyuk et al. 2020). The HOOK complex promotes invasion by regulating microneme exocytosis To determine whether the HOOK complex is required for microneme exocytosis, we directly measured secretion of the microneme protein MIC2. Microneme exocytosis exposes integral membrane proteins like MIC2 that function as adhesins during gliding motility. These adhesins subsequently undergo proteolytic cleavage and are released into the supernatant, which can be analyzed by immunoblot (V. B. Carruthers, Sherman, and Sibley 2000). Quantification of the proportion of secreted MIC2 was performed by generating a standard curve from unstimulated lysates. Strikingly, parasites depleted of either member of the HOOK complex were severely impaired in their ability to secrete microneme proteins (Figure 9J, Figure 10G). Inhibition of the complex led to a similar degree of secretion impairment as CDPK1 depletion. Together, these data suggest that HOOK and FTS form a functional complex that is required for microneme exocytosis. 75 Figure 10. Extended analysis of FTS knockdown, proximity labeling, and microneme protein secretion. (A) PCR analysis confirming correct integration of the mAID-HA cassette at the C terminus of endogenous FTS (TGGT1_264050) in the TIR1 line. (B) Uncropped immunoblot shown in Figure 5D confirming C-terminal tagging of FTS. The band at ~70 kDa represents anti-HA/anti-ALD-related background present in all conditions. © Replication assays of host cells infected with TIR1 or FTS-AID parasites in auxin for 24 hrs. Parasites per vacuole were quantified from immunofluorescence on fixed intracellular parasites. p > 0.9. Two-way ANOVA. (D) Live microscopy of HFFs infected with parasites expressing cytosolic mNeonGreen-TurboID as the cytosolic control for proximity labeling. (E) PCR analysis confirming presence of TurboID-Ty cassette in the TIR1 line. (F) Immunoblot detection of biotinylated proteins in FTS-TurboID and cytosolic mNG-TurboID parasites treated with 500 µM of biotin or a vehicle of DMSO. Biotinylated proteins detected with a labeled streptavidin. anti-CDPK1 antibody was used as a loading control. (G) Serial dilution of total parasite lysate for cKD strains for TIR1, CDPK1, HOOK, and FTS used in microneme protein secretion assays in Figure 5J to generate standard curves. 76 Microneme trafficking depends on CDPK1 activity and HOOK We hypothesized that if the HOOK complex is required for sustained microneme exocytosis, then depletion of HOOK would inhibit trafficking of micronemes during parasite motility. Dynamic relocalization of micronemes can be visualized using live microscopy of parasites expressing the integral microneme protein CLAMP fused to mNeonGreen (CLAMP-mNG) (Figure 11A)(Sidik, Huet, et al. 2016b). To trigger microneme exocytosis, parasites are treated with zaprinast to stimulate a rise in intracellular Ca2+ and activate CDPK1. Dynamic relocalization of micronemes is defined as the CLAMP-mNG signal that concentrates at the apical end of the parasite over time. To quantify microneme relocalization, mNG signal is measured, over time, along the apical-basal axis of individual parasites within a vacuole. We scored microneme relocalization by calculating the difference in maximum apical intensity comparing the instant of zaprinast addition to the time point just prior to parasite egress of the uninhibited zaprinast-stimulated controls (Figure 11A). We first assessed whether CDPK1 kinase activity is required for microneme relocalization using the specific inhibitor 3-MB-PP1 (Lourido et al. 2010). We incubated parasites with 3-MB-PP1 for 30 min, imaged for 1 min to establish a baseline, and then stimulated with zaprinast. Parasites pre-treated with DMSO relocalized micronemes as expected; however, microneme relocalization was blocked when CDPK1 was inhibited by 3-MB-PP1 (Figure 11B–D). These results indicate microneme relocalization depends on CDPK1 kinase activity. We next assessed whether HOOK is also required for microneme relocalization. We fused an mNG reporter to the C terminus of CLAMP in AID-HOOK parasites and TIR1 parasites as a control (Figure 12A–B). We treated parasites with auxin for 24 hrs prior to stimulation with zaprinast. Microneme relocalization was unaffected in TIR1/CLAMP-mNG parasites treated with auxin (Figure 11E–F). By contrast, microneme relocalization did not occur in intracellular AID-HOOK parasites regardless of auxin treatment (Figure 11G–H). We attributed the lack of relocalization in vehicle-treated parasites to the hypomorphism resulting from N-terminal tagging, which is consistent with the delayed invasion and reduced plaque size documented for this strain. To 77 determine whether vehicle-treated AID-HOOK parasites relocalized micronemes over longer periods, we examined CLAMP-mNG localization in extracellular parasites after egress. While vehicle-treated AID-HOOK parasites were able to relocalize micronemes, parasites depleted of HOOK exhibited a striking microneme localization dispersed throughout the cytosol (Figure 11I–J). As expected, TIR1/CLAMP-mNG parasites maintained the apical relocalization of micronemes regardless of auxin treatment in extracellular parasites. We used an alternative metric for microneme relocalization to account for the disorganized localization of micronemes in parasites depleted of HOOK. We quantified the percent of the total CLAMP-mNG signal localized to the apical end versus the remaining body of the parasite (Figure 11K). The apical region cutoffs were experimentally derived from vehicle-treated TIR1/CLAMP-mNG parasites and defined as the apical 12.5% of the parasite. While a majority of CLAMP-mNG signal was localized to the apical region of TIR1/CLAMP-mNG parasites, most of the CLAMP-mNG signal observed in parasites depleted of HOOK was found in the parasite body. Micronemes rapidly adopt an aberrant localization in the absence of HOOK during parasite motility. These results indicate that CDPK1 kinase activity and HOOK are required for microneme trafficking during parasite motility. 78 Figure 11. CDPK1 activity and HOOK are required for microneme trafficking during parasite motility stages. (A) Schematic to analyze microneme trafficking during parasite motile stages. Intracellular parasites expressing microneme protein CLAMP endogenously tagged with mNeonGreen (CLAMP-mNG). Parasites are treated either with 3-MB-PP1 (inhibit CDPK1) or auxin (for conditional knockdown). Live microscopy was performed to detect CLAMP-mNG signal over time. Zaprinast was added at 1 min or 30 sec to stimulate microneme relocalization to the apical end of the parasite. Fluorescence intensities across the apical-basal axis of each individual parasite within a vacuole was measured across time. Microneme relocalization was quantified by calculating the difference of maximum CLAMP intensity between time points preceding drug addition and egress. (B) Maximum intensity projections at single time points of CLAMP-mNG parasites treated with 3 µM 3-MB- PP1 or vehicle and zaprinast. (C) Relative fluorescence intensity of CLAMP-mNG signal across the apical-basal axis of parasites in B. Zaprinast (red) or vehicle (blue). Splines mean intensity for all parasites in each vacuole are shown with SD shaded. (D) Microneme relocalization. SuperPlots showing vacuole median peak differences are displayed as triangles. Individual parasites are displayed as circles. Replicates are differentially shaded, n.s., p > 0.05, unpaired t-test. (E) Maximum intensity projections at single time points of TIR1/CLAMP-mNG parasites treated with auxin and stimulated with zaprinast. (F) Microneme relocalization was quantified for TIR1/CLAMP-mNG parasites as in D. (G) Maximum intensity projections at single time points of AID-HOOK/CLAMP-mNG parasites treated auxin and stimulated with zaprinast. (H) Microneme relocalization was quantified for AID-HOOK/CLAMP-mNG parasites as in D. (I) Maximum intensity projections of extracellular TIR1/CLAMP-mNG and AID-HOOK/CLAMP-mNG parasites. (J) Percent of extracellular parasites in I with WT CLAMP-mNG localization, n.s., p > 0.05, Welch’s t-test. (K) Percent total CLAMP-mNG signal intensity in the apical versus body of extracellular parasites, n.s., p > 0.05, Welch’s t-test. 79 Figure 12. Extended analysis of FTS knockdown, proximity labeling, and microneme protein secretion. (A) PCR analysis confirming correct integration of the mNeonGreen reporter at the C terminus of endogenous CLAMP (TGGT1_265790) in the TIR1 line. (B) PCR analysis confirming correct integration of the mNeonGreen reporter at the C terminus of endogenous CLAMP in the AID-HOOK line. DISCUSSION In this study, we characterized a new regulator of microneme exocytosis called HOOK, which forms a stable complex with FTS and two other proteins. Homologs of HOOK and FTS participate in dynein-mediated vesicular trafficking in other organisms. In T. gondii, knockdown of HOOK or FTS blocked invasion of host cells and altered rapid microneme trafficking during Ca2+-regulated motility. Overall, we show how studying parasite signaling pathways can illuminate the cellular adaptations that support parasitism. Although CDPK1 is known to mediate microneme exocytosis (Lourido et al. 2010), the precise molecular events controlling this process have remained elusive. In this study, we determined that CDPK1 activity is required for the Ca2+-stimulated trafficking of micronemes to the apical end. This phenotype depends on HOOK, one of the CDPK1 targets uncovered by both global phosphoproteomics and thiophosphorylation studies. Despite prior annotation as a microneme protein (Butler et al. 2014), HOOK likely functions as an activating adaptor in T. gondii based on homology to other Hook proteins and the functional characterization we have performed. In particular, immunoprecipitation of HOOK identified three interacting proteins: FTS, TGGT_316650, and TGGT1_306920. In H. sapiens, A. nidulans, and D. melanogaster, Hook proteins 80 complex with FTS and FHIP (the FHF complex), to link cargo to the dynein machinery for trafficking along microtubules (Bielska et al. 2014; Xu et al. 2008; Yao, Wang, and Xiang 2014; Gillingham et al. 2014; Guo et al. 2016). FHF complexes have been shown to recognize early endosomes via FHIP binding to RAB5 (Bielska et al. 2014; Zhang et al. 2014; Guo et al. 2016). Homology to FHIP was lacking in TGGT1_316650 and TGGT1_306920. However, genome-wide knockout screens indicate TGGT1_306920 is critical for parasite fitness, leading us to speculate that it may mediate cargo binding in a manner structurally distinct from known adaptors. Additional work will be needed to elucidate the function of the HOOK complex and to understand how it mediates trafficking of apicomplexan-specific organelles. Our results place the HOOK complex downstream of CDPK1, regulating the stimulated relocalization of micronemes to the apical end of parasites. However, the HOOK complex only partially explains CDPK1’s role during egress. Whereas CDPK1 depletion blocks egress entirely, parasites depleted of HOOK or FTS exit host cells, despite subsequent defects in microneme secretion and invasion. Microneme discharge is required for egress, supplying adhesins for gliding motility and the perforin PLP1 to rupture the parasitophorous vacuole membrane (Kafsack et al. 2009). It therefore appears that the initial round of microneme discharge during egress depends on CDPK1, and only subsequent rounds require the HOOK complex. Indeed, a fraction of micronemes are already found docked at the apical complex prior to the transition from the replicative to the motile stages, and may constitute the first round of microneme exocytosis (Sun et al. 2022; Mageswaran et al. 2021). Dynein-mediated trafficking might become important immediately following egress when continuous microneme discharge is necessary, as is the case during gliding motility and invasion. Consistent with this hypothesis, depletion of the dynein light chain 8a (DLC8a) yielded similar phenotypes to HOOK and FTS knockdown: intact egress but dysfunctional microneme protein secretion, gliding motility, and invasion (Lentini et al. 2019). Loss of DLC8a also led to defects in rhoptry positioning, which were not observed in HOOK knockdowns (Lentini et al. 2019), so other activating adaptors may be involved in rhoptry trafficking. Our 81 results suggest that the HOOK complex has been specifically adapted for microneme trafficking in T. gondii. Several aspects of microneme trafficking remain to be determined. The precise nature of the dynein complex is poorly understood. Eukaryotes typically express a single cytoplasmic dynein heavy chain. In T. gondii, TGGT1_294550 is the top candidate for the cytoplasmic dynein heavy chain (DHC) as it contains the necessary domains, is conserved among apicomplexans, and is required for parasite fitness (Lentini et al. 2019; Sidik, Huet, et al. 2016b). Other putative DHCs in the T. gondii genome are likely axonemal dyneins required for flagellar function in sexual-stage parasites. It also remains unknown how the HOOK complex binds to micronemes. In H. sapiens and D. melanogaster, RAB5 on vesicles interacts with FHIP in the HOOK complex (Bielska et al. 2014; Xu et al. 2008; Yao, Wang, and Xiang 2014; Gillingham et al. 2014; Guo et al. 2016). We speculate that TGGT1_306920 may serve the role of FHIP within the HOOK complex but its binding partner on micronemes remains unknown. RAB5A and RAB5C have been implicated in the biogenesis of micronemes, but their roles during exocytosis have not been explored (Kremer et al. 2013). Understanding how micronemes are recognized may elucidate how cargo specificity is achieved and regulated. Ca2+ signaling has been tuned to support apicomplexan-specific cellular processes but identifying and integrating effectors within the pathway has remained a major bottleneck. Signaling effectors conserved amongst eukaryotes are often integrated into apicomplexan cellular pathways in novel or unusual ways. Here, we identified candidate downstream effectors of the Ca2+-regulated kinase CDPK1 by monitoring proteome-wide protein phosphorylation. In addition to creating a catalog of candidates for future characterization, we identified, characterized, and integrated the HOOK complex downstream of CDPK1 within the Ca2+ signaling network controlling microneme exocytosis. Lastly, our results are the first to implicate activating adaptors as critical factors for the pathogenesis of an apicomplexan organism by promoting exocytic trafficking. 82 MATERIALS & METHODS Cell culture T. gondii parasites were grown in human foreskin fibroblasts (HFFs, ATCC SRC-1041) maintained in DMEM (GIBCO 11965118) supplemented with 3% inactivated fetal calf serum and 10 µg/mL gentamicin (Thermo Fisher Scientific), referred to as media. When noted, DMEM was supplemented with 10% inactivated fetal bovine serum (IFS) and 10 µg/mL gentamicin, referred to as 10% IFS media. Parasites and HFFs were grown at 37°C/5% CO2 unless indicated otherwise. Parasite transfection and strain construction Genetic background of parasite strains T. gondii RH strains were used as genetic backgrounds for this study. All strains contain the ∆ku80∆hxgprt mutations to facilitate homologous recombination (Huynh and Carruthers 2009). TIR1 expresses the TIR1-FLAG ubiquitin ligase and the CAT enzyme conferring chloramphenicol resistance (Brown, Long, and Sibley 2017). Transfection Parasites were pelleted at 1000 x g for 5 to 10 min and resuspended with Cytomix (10 mM KPO4, 120 mM KCl, 0.15 mM CaCl2, 5 mM MgCl2, 25 mM HEPES, 2 mM EDTA, 2 mM ATP, and 5 mM glutathione) and combined with DNA to a final volume of 400 µL. Parasites were electroporated using an ECM 830 Square Wave electroporator (BTX) in 4 mm cuvettes with the following setting: 1.7 kV, 2 pulses, 176 µs pulse length, and 100 msec interval. Endogenous tagging of genes Genes were endogenously tagged using the previously described high-throughput tagging (HiT) strategy (Smith et al. 2022). Cutting units specific to each gene were purchased as gene fragments (IDT gBlocks; P22 and P23) and integrated with the following empty HiT vector backbones via Gibson assembly: pALH086 (V5-mAID-HA; GenBank: ON312869), pALH047 (V5-3HA; GenBank: ON312868), and pALH173 (TurboID-Ty; GenBank: Pending). Between 30 and 50 µg of each vector was linearized with BsaI and co-transfected with 50 µg of the pSS014 Cas9 expression plasmid (GenBank: OM640002). Vectors targeting TGGT1_289100 (HOOK) were transfected into TIR1 or CDPK1-AID. Vectors targeting TGGT1_264050 (FTS) were transfected into TIR1. Parasites underwent drug selection for approximately 1 week in 10% IFS media with 3 µM pyrimethamine or 25 µg/mL mycophenolic acid and 50 µg/mL xanthine, followed by subcloning into 96-well plates. Single clones were screened for tag expression by PCR, immunofluorescence, or immunoblot. PCR validation was performed using the primers P44/P45 (FTS-AID) and P46/P47 (FTS-TurboID). Endogenous tagging of HOOK N terminus 83 AID-HOOK N-terminal tagging was generated by PCR amplifying the gene fragment encoding the HA-mAID with homology arms to HOOK (IDT gBlock; P17) using the primers P18/19. A sgRNA targeting HOOK was assembled into the pSS013 Cas9 expression plasmid (GenBank: OM640003) using the primers P20/P21. 10 µg of the PCR product was transfected with 50 µg of the Cas9 expression plasmid. Parasites were subcloned into 96-well plates. Single clones were screened using PCR primers P42/P43 and validated by immunofluorescence and immunoblot. TIR1/pMIC2-mNeonGreen-TurboID-Ty pMIC2-mNG-TurboID-Ty was amplified with primers P24/P25 from pALH184 (pMIC2- mNG-TurboID-Ty-3'DHFR; GenBank: Pending) with homology arms to the 5′ and 3′ ends of a defined, neutral genomic locus (Markus et al. 2019). Amplified DNA was co- transfected with the Cas9 expression plasmid targeting the neutral locus (pBM041; GenBank: MN019116). mNeonGreen-expressing parasites were isolated by FACS and subcloned in 96-well plates. Expression was confirmed by fluorescence. Endogenous tagging of CLAMP mNeonGreen DNA was amplified using PCR from pGL015 (V5-mNG-mAID-Ty; GenBank: OM640005) with homology arms targeting the C terminus of TGGT1_265790 (CLAMP) using the primers P26/P27. A sgRNA targeting CLAMP was assembled into the pSS013 Cas9 expression plasmid (GenBank: OM640003) using the primers P28/P29. Between 5 and 10 µg of mNeonGreen DNA was co-transfected with 50 µg of the Cas9 expression plasmid targeting CLAMP into TIR1 and AID-HOOK. Parasites were grown in 10% IFS media until lysing the HFF monolayer, followed by FACS-mediated isolation of mNeonGreen expressing parasites and subcloning into 96-well plates. Single clones were screened for mNeonGreen expression by PCR using primers P48/P49 and live microscopy. Immunoblotting Samples were combined with 5X Laemmli sample buffer (10% SDS, 50% glycerol, 300mM Tris HCl pH 6.8, 0.05% bromophenol blue, 5% beta-mercaptoethanol) and were incubated at 95°C for 10 min. The samples were run on precast 4–15% or 7.5% SDS gels (Bio-Rad) and were transferred overnight onto a nitrocellulose membrane in transfer buffer (25mM TrisHCl, 192 mM glycine, 0.1% SDS, 20% methanol) at 4°C. Blocking was performed with 5% milk in PBS for 1 hr rocking at room temperature. Antibody incubations were performed with 5% milk in TBS-T for 1 hr rocking at room temperature. Three 5 min TBS-T washes were performed before and after secondary antibody incubations rocking at room temperature. After a final PBS wash, imaging was performed using a LI-COR Odyssey. For immunoblot detection of the HA tag in AID-HOOK and FTS-AID after auxin-mediated depletion, secondary antibody incubations were performed with anti-rabbit HRP antibodies (Jackson ImmunoResearch) and detected with chemiluminescence (Azure) for increased sensitivity. Imaging was performed using a Bio-Rad Gel Doc XR. 84 Immunofluorescence analysis HFFs were seeded onto coverslips and grown until confluence. Confluent HFFs were infected with parasites. Approximately 2 hrs later, media was exchanged with media containing either 50 µM auxin or vehicle solution of PBS. At 24 hrs post-infection, the media was aspirated, and the coverslips were washed with PBS three times before fixation with 4% formaldehyde in PBS for 20 min. Following three washes in PBS, the fixed cells were permeabilized with 0.25% Triton X-100 for 15 min. After three washes in PBS, the coverslips were incubated in a blocking solution (5% IFS/ 5% NGS in PBS) for 10 min at room temperature. Coverslips were incubated for 1–2 hrs with primary antibody diluted in blocking solution. Anti-GAP45 or anti-MIC2 was used as a parasite counterstain. After three washes in PBS, the coverslips were incubated in blocking solution for 5 min, followed by secondary antibody diluted in blocking solution containing Hoechst 33342 for 1 hr. The coverslips were washed three times in PBS, once in water, and finally mounted with Prolong Diamond overnight at room temperature. Microscope images were acquired with the Nikon Ti Eclipse and NIS Elements software package. Plaque Assays 600 and 1200 parasites were used to infect 6-well plates of HFFs in 10% IFS media. At 1 dpi, media was exchanged with 10% IFS media containing either 50 µM auxin or vehicle solution of PBS. Parasites were allowed to grow undisturbed for 8 days total. Plates were washed with PBS and fixed for 10 min with 100% ethanol at room temperature. After removing ethanol, plates were allowed to dry prior to staining with crystal violet solution for 30 min to 1 hr. Plates were washed 3 times with PBS, once with water, and allowed to dry before scanning. Invasion assays Confluent HFFs seeded in T12.5 flasks were infected with parasites. Approximately 2 hrs later, the media was exchanged for 10% IFS media containing either 50 µM auxin or vehicle solution of PBS. Parasites were harvested at 2 dpi and the media was exchanged for 1% IFS in invasion media (DMEM Sigma D2902, 20 mM HEPES, pH 7.4) with auxin or vehicle PBS added. Confluent HFFs seeded in clear-bottomed 96-well plates were infected with 2 x 105 extracellular parasites. After centrifuging the plate at 290 x g for 5 min to synchronize invasion, the infected 96-well plate was incubated by floating on a water bath at 37°C/5% CO2 for 90 min. Wells were washed once with PBS before fixing infected HFFs with 4% formaldehyde in PBS for 20 min at room temperature. Wells were washed three times with a wash buffer (1% NGS in PBS) and then incubated in a blocking buffer (5% IFS and 1% NGS in PBS) overnight at 4°C. To stain extracellular parasites, wells were incubated with anti-SAG1 antibody for 30 min at room temperature. After washing wells three times with a wash buffer, fixed HFFs were permeabilized in 0.25% TritonX-100 in blocking buffer for 8 min room temperature. After washing the wells three times with a wash buffer, the wells were incubated in a blocking buffer for 10 min at room temperature, followed by anti-GAP45 antibody for 30 min at room 85 temperature to label all parasites. After three washes in a wash buffer, wells were incubated with a secondary antibody solution containing Hoechst for 30 min. After three washes in PBS, wells were imaged using a Biotek Cytation 3. Egress assays Confluent HFFs seeded in glass bottomed 35 mm dishes (Ibidi or Mattek) were infected with approximately 2 x 105 parasites. Approximately 2 hrs later, the media was exchanged for 10% IFS media containing either 50 µM auxin or vehicle solution of PBS. At 24 hrs post-infection, the media was replaced with 1% IFS in Ringer’s (155 mM NaCl, 2 mM CaCl2, 3 mM KCl, 1 mM MgCl2, 3 mM NaH2PO4, 10 mM HEPES, 10 mM glucose, pH 7.4). Zaprinast (final concentration 500 µM) was added into the dishes after 15 sec of imaging and imaging of infected HFFs continued for a total of 10 min. Images were acquired using a Nikon Ti Eclipse with an enclosure maintained at 37°C. The number of intracellular vacuoles was quantified at 0 min and 10 min. Results are the sum of four fields of view per condition and are representative of three independent experiments. Replication assays Confluent HFFs seeded on coverslips were infected with parasites. Parasites were centrifuged at 290 x g for 5 min to synchronize invasion of host cells. Approximately 2 hrs later, the media was exchanged for 10% IFS media containing either 50 µM auxin or vehicle solution of PBS. At 24 hrs post-infection, the media was aspirated and fixed as described in “Immunofluorescence Analysis”. Coverslips were incubated in primary and secondary antibodies for 30 min. Anti-GAP45 antibody was used to visualize parasites. Microscope images were acquired with the Nikon Ti Eclipse and NIS Elements software package. Tiled images were acquired in a four-by-four manner using a 40x objective. Parasites per vacuole were quantified for at least 100 vacuoles for three biological replicates. Immunoprecipitation of HOOK and FTS complex Parasite harvest HOOK-3xHA, CDPK1-AID (parental), FTS-3xHA and TIR1 (parental) parasites were infected onto confluent HFFs in 15 cm dishes. After the parasites lysed the HFF monolayer (approximately 40 hrs post-infection), extracellular parasites were passed through 5 µm filters and washed twice with DMEM by pelleting at 1000 x g for 10 min. Parasite pellets were resuspended in 1X Dynein Lysis Buffer (DLB) (30 mM HEPES, 50 mM KOAc1, 2 mM MgOAc, 10% glycerol, 1 mM EGTA pH 8.0, 1 mM DTT, 0.5 mM ATP, 125 units/mL benzonase, 1X HALT protease and phosphatase inhibitor, and 1% NP-40 IGEPAL CA 630) to achieve a concentration of 6.67 x 108 parasites/mL (Redwine et al. 2017). Parasites were lysed on ice for 10 min and vortexed periodically to facilitate lysis. Lysates were centrifuged at 1000 x g for 5 min at room temperature to pellet unlysed parasites and the lysate supernatant was collected and used as the immunoprecipitation input. 86 Immunoprecipitation 25 µL of anti-HA magnetic beads (Thermo Scientific 88836) was used for 200 µL of parasite lysate. Beads were aliquoted and washed three times with a wash buffer (30 mM HEPES, 50 mM KOAc1, 2 mM MgOAc, 10% glycerol, 1 mM EGTA pH 8.0, 1 mM DTT, 0.5 mM ATP, and 0.01% NP-40 IGEPAL CA 630) using a magnetic rack. To begin the pulldown, 200 µL of parasite lysate was used to resuspend washed beads. Tubes were rotated at 4°C for 1 hr (FTS-AID and TIR1) and 3 hrs (HOOK-3xHA and CDPK1- AID). Beads were resuspended in 250 µL of wash buffer, transferred to a new tube, and received two additional washes with the same volume. Proteins were eluted by resuspending beads in 22 µL of 1X S-trap sample buffer (5% SDS, 50 mM TEAB, pH 7.5) and incubated at 70°C for 10 min. The eluate was collected for MS sample processing and analysis. Results are representative of three independent experiments. Protein cleanup and digestion Proteins were prepared for mass spectrometry as described above in “Sub-minute phosphoproteomics - Protein cleanup and digestion”. Eluted peptides were frozen in liquid nitrogen, lyophilized, and stored at -80°C until MS analysis. MS data acquisition Lyophilized peptides were resuspended in 25 µL of 0.1% formic acid and were analyzed on an Exploris 480 Orbitrap mass spectrometer equipped with a FAIMS Pro source (Bekker-Jensen et al. 2020) connected to an EASY-nLC chromatography system using 0.1% formic acid as Buffer A and 80% acetonitrile/0.1% formic acid as Buffer B. Peptides were separated at 300 nL/min on a gradient of 1–6% B for 1 min, 6–21% B for 41 min and 30 sec, 21–36% B for 20 min and 45 sec, 36–50% B for 10 min and 15 sec, 100% B for 14 min and 30 sec, 100–2% B for 3 min, 2% B for 3 min, 2–98% B for 3 min, and 98% B for 3 min. The orbitrap and FAIMS were operated in positive ion mode with a positive ion voltage of 1800V; with an ion transfer tube temperature of 270°C; using a standard FAIMS resolution and compensation voltage of -50 and -65V, an inner and outer electrode temperature of 100°C with 4.5 mL/min carrier gas. Full scan spectra were acquired in profile mode at a resolution of 60,000, with a scan range of 350-1400 m/z, 300% AGC target, 25 msec maximum injection time, intensity threshold of 5 x 103, 2–6 charge state, dynamic exclusion of 20 sec, 15 data dependent scans (DDA Top 15), and mass tolerance of 10 ppm. MS2 spectra were generated an HCD collision energy of 30 at a resolution of 15,000, first mass at 110 m/z, with an isolation window of 1.3 m/z, and an automatically determined AGC target and maximum injection time in standard and auto mode. Immunoprecipitation MS analysis Raw files were analyzed in Proteome Discoverer 2.4 (Thermo Fisher Scientific) to generate peak lists and protein and peptides identifications using Sequest HT (Thermo Fisher Scientific) and the ToxoDB release 49 GT1 protein database. The maximum missed cleavage sites for trypsin was limited to 2. The following modifications were included in the search: dynamic oxidation (+15.995 Da; M), dynamic phosphorylation (+79.966 Da; S,T,Y), dynamic acetylation (+42.011 Da; N-terminus), and static methylthio 87 (+45.988 Da; C). Label free quantification of proteins was performed using summed abundances from unique peptides. Abundances were normalized on the total peptide amount. Pairwise ratios were calculated for protein abundances comparing strains and conditions. Significance values were derived from t-tests across three replicates and adjusted with Benjamini-Hochberg correction. Exported protein abundance files from Proteome Discoverer 2.4 were loaded into R (version 4.1.1). Proximity labeling FTS-TurboID Parasite harvest and treatment FTS-TurboID and mNG-TurboID (cytosolic control) parasites were infected onto confluent HFFs in 15 cm dishes. After the parasites lysed the HFF monolayer (approximately 40 hrs post-infection), extracellular parasites were passed through 5 µm filters and washed twice with DMEM by pelleting at 1000 x g for 10 min. At least 1 x 108 Parasites were resuspended in 200 µL of DMEM. 200 µL of biotin (final concentration 500 µM) or vehicle DMSO in DMEM was mixed with parasites. Tubes were incubated in a water bath at 37°C for 5 min. Parasites were pelleted at 12,000 x g for 1 min at 4°C for a total of three 1 mL PBS washes. Parasites were resuspended in a RIPA NP-40 lysis buffer (10 mM Tris-HCl pH 7.5, 140 mM NaCl, 1% NP-40 IGEPAL CA 630, 0.1% sodium deoxycholate, 0.1% SDS, 1x HALT protease inhibitor, and 125 units/mL benzonase) at a parasite concentration of 5 x 108 parasites/mL. Parasites were lysed at room temperature for 15 min and stored at -20°C. Results are representative of three independent experiments. Biotinylated protein enrichment Streptavidin magnetic beads were washed three times with a RIPA NP-40 lysis buffer. Thawed lysates were spun at 16,000 x g for 5 min at 4°C and the supernatant was used as the pulldown input. 20 µL of streptavidin magnetic beads was used for 200 µL of lysates (1 x 108 parasites). Beads were incubated with lysate rotating for 1 hr. Beads received a series of 1 mL washes: twice with the RIPA NP-40 lysis buffer, once with 1 M KCl, once with 0.1M Na2CO3 (pH 11), once with 2 M urea in 10 mM Tris pH 8.0), and twice with RIPA NP-40 lysis buffer. Proteins were eluted by incubating beads with 2.5 mM biotin in S-trap sample buffer (5% SDS, 50 mM TEAB, pH 7.5) for 10 min at 95°C. Protein cleanup and digestion Proteins were prepared for mass spectrometry as described above in “Sub-minute phosphoproteomics - Protein cleanup and digestion”. Eluted peptides were frozen in liquid nitrogen, lyophilized, and stored at -80°C until MS analysis. MS data acquisition Lyophilized peptides were resuspended in 25 µL of 0.1% formic acid and were analyzed on an Exploris 480 Orbitrap mass spectrometer equipped with a FAIMS Pro source (Bekker-Jensen et al. 2020) connected to an EASY-nLC chromatography system using 0.1% formic acid as Buffer A and 80% acetonitrile/0.1% formic acid as Buffer B. Peptides were separated at 300 nL/min on a gradient of 2% B for 1 min, 2–25% B for 41 88 min, 25–40% B for 6 min, 40–100% B for 12 min, 100–2% B for 3 min, 2% B for 3 min, 2–98% B for 3 min, and 98% B for 3 min. The orbitrap and FAIMS were operated in positive ion mode with a positive ion voltage of 1800V; with an ion transfer tube temperature of 270°C; using a standard FAIMS resolution and compensation voltage of -50 and -65V, an inner and outer electrode temperature of 100°C with 4.5 mL/min carrier gas. Full scan spectra were acquired in profile mode at a resolution of 60,000, with a scan range of 350–1400 m/z, 300% AGC target, 25 msec maximum injection time, intensity threshold of 5 x 103, 2–6 charge state, dynamic exclusion of 20 sec, 15 data dependent scans (DDA Top 15), and mass tolerance of 10 ppm. MS2 spectra were generated with a HCD collision energy of 30 at a resolution of 15,000, first mass at 110 m/z, with an isolation window of 1.3 m/z, and a normalized AGC target of 200% with an automatically determined maximum injection time. Proximity labeling MS analysis Raw files were analyzed in Proteome Discoverer 2.4 (Thermo Fisher Scientific) to generate peak lists and protein and peptides identifications using Sequest HT (Thermo Fisher Scientific) and the ToxoDB release 49 GT1 protein database. The maximum missed cleavage sites for trypsin was limited to 2. The following modifications were included in the search: dynamic oxidation (+15.995 Da; M), dynamic phosphorylation (+79.966 Da; S,T,Y), dynamic biotinylation (+226.078 Da; K, N-terminus), dynamic acetylation (+42.011 Da; N-terminus), and static methylthio (+45.988 Da; C). Label free quantification of proteins was performed using summed abundances from unique peptides. Abundances were normalized on the total peptide amount. Pairwise ratios were calculated for protein abundances comparing strains and conditions. Significance values were derived from t-tests across three replicates and adjusted with Benjamini- Hochberg correction. Exported protein abundance files from Proteome Discoverer 2.4 were loaded into R (version 4.1.1). MIC2 secretion assays Parasite harvest and treatment Extracellular parasites were harvested in chilled DMEM and washed twice in DMEM after centrifugation at 1000 x g for 10 min at 4°C. Parasites were resuspended at a concentration of 6 x 108 parasites/mL in cold DMEM. 3 x 107 parasites were aliquoted into round-bottom 96-well plates. An additional aliquot of parasites was lysed in 5X Laemmli sample buffer (see “Immunoblotting” for recipe) to obtain the total parasite lysate used to determine total MIC2 levels. Secretion was stimulated in plates with IFS and ethanol in DMEM (final concentration 3% IFS and 1% ethanol) or a vehicle solution of DMEM. Plates were incubated by floating on a water bath at 37°C/5% CO2 for 90 min. Plates were spun at 1000 x g for 5 min at 4°C to separate parasites from secreted proteins. Supernatants were transferred to a new well and spun again. The final supernatant was mixed with a 5X Laemmli sample buffer, boiled at 95°C for 10 min, and stored at -20°C along with total lysates until immunoblot analysis. Immunoblot analysis and quantification 89 To quantify MIC2 protein levels, a standard curve derived from the total parasite lysate was generated alongside supernatants containing secreted microneme proteins. The standard curve was derived from 3-fold serial dilutions of the total parasite lysate (undiluted, 1:3, 1:9, and 1:27) in a 1X Laemmli sample buffer. The serial dilutions and supernatants of auxin- and vehicle-treated parasites of a single strain were loaded onto the same precast 4–15% gel (Bio-Rad). Subsequent immunoblotting steps were performed as described in “Immunoblotting”. Anti-MIC2 was used to detect total and secreted MIC2 proteins. Secreted MIC2 proteins have a lower molecular weight due to parasite-mediated proteolytic cleavage. Anti-CDPK1 was used as a loading control for total parasite lysate and to reveal any parasite lysis or carry over in the supernatant. Immunoblots confirming the depletion of tagged proteins were also collected. Imaging was performed on a LI-COR Odyssey at high resolution for quantification. Immunoblot quantification was performed in Fiji on unadjusted inverted images. For a single immunoblot, lane profiles of uniform dimensions were generated for MIC2 signal. Background-subtracted signal intensity was measured as an area using the line and wand tool. A standard curve was derived from the quantified MIC2 signal of the dilution series. Standard curves across all conditions and replicates had R2 values greater than 0.90. Secreted MIC2 signals were within the linear range of the standard curve and were used to calculate the percent of total MIC2 secreted. Results are representative of three independent experiments for each parasite strain. Microneme relocalization Parasites expressing the CLAMP-mNG reporter were grown in HFFs in glass-bottomed 35mm dishes (Ibidi and Mattek) for approximately 20 hrs. For 3-MB-PP1 treatment, media was exchanged 30 min prior to live microscopy for 3% IFS in Ringer’s buffer (155 mM NaCl, 2mM CaCl2, 3mM KCl, 1mM MgCl2, 3mM NaH2PO4, 10mM HEPES, 10 mM glucose, pH 7.4) containing either 3 µM 3-MB-PP1 or vehicle solution of DMSO. At approximately 1 min after beginning live microscopy, parasites were stimulated with zaprinast (500 µM final concentration) or a vehicle solution of DMSO in corresponding Ringer’s buffer. For TIR1/CLAMP-mNG and AID-HOOK/CLAMP-mNG parasite, media was exchanged 2 hrs after infection for 10% IFS media containing either 50 µM auxin or vehicle solution of PBS. Media was exchanged for 3% IFS in Ringer’s buffer just prior to live microscopy. At approximately 30 sec after beginning live microscopy, parasites were stimulated with zaprinast (500 µM final concentration) or a vehicle solution of DMSO in corresponding Ringer’s buffer. Images were recorded every 5–7 sec until egress or approximately 5 min using a Nikon Ti Eclipse with an enclosure maintained at 37°C. Microneme relocalization was quantified using built-in commands from ImageJ (v. 1.53e). To determine the distribution of fluorescent micronemes at a specific time frame, a line was drawn from the parasite’s apical end to its basal end. The “plot profile” command was applied to this line to calculate fluorescence intensity at regularly spaced intervals (0.13 microns) across the parasite on unadjusted images. This process was repeated for each time frame of interest and for each parasite in the analyzed vacuole. 90 Regression analysis was performed on relocalization data from each time frame, resulting in logarithmic regression plots that display how fluorescence intensity correlates with distance from the parasite’s apical end. Microneme relocalization is shown as SuperPlots to simultaneously visualize the median relocalization of an entire vacuole and the relocalization of individual parasites within each vacuole (Lord et al. 2020). Microneme localization in extracellular parasites Microneme localization in extracellular parasites was quantified using a custom-built, image analysis pipeline. ImageJ’s built-in commands were used to measure the length of the parasite’s major axis. Based on the patterns of vehicle-treated TIR1/CLAMP-mNG data, 1/8th of the parasite closest to the apical end was deemed the apical region. The remaining region was deemed the body. Using the Interactive Learning and Segmentation Toolkit (ilastik v. 1.3.3) (Berg et al. 2019), the apical region and body of parasites were isolated from each other and from the surrounding media. Ilastik’s pixel and object quantification tools were then used to determine the total intensity of fluorescence in each of these regions. Comparing these values to each other yielded the percentage of fluorescent micronemes present in each of these sections in relation to the total fluorescence present in the parasite. This process was repeated for several extracellular parasites in each of the conditions tested. 91 AUTHOR CONTRIBUTIONS Alex W Chan Contribution: Conceptualization, Methodology, Validation, Formal analysis, Investigation, Writing - original draft, Writing - review and editing, Visualization Nicole Haseley Contribution: Methodology, Formal analysis Sundeep Chakladar Contribution: Methodology, Formal analysis, Investigation Elena Andree Contribution: Investigation Alice L Herneisen Contribution: Resources, Methodology Sebastian Lourido Contribution: Conceptualization, Resources, Supervision, Funding acquisition, Methodology, Writing - review and editing 92 REAGENT AND FUNDING RECOGNITION We thank Forest M. White for generous support in developing the mass spectrometry methodologies used in this study and for helpful discussions. For data in Figure 2, we thank Matthew Child and Matt Bogyo for the CDPK1 antibody, Peter Bradley for the GAP45 antibody, John Boothroyd for the SAG1 antibody. We thank Marc-Jan Gubbels for the alpha-tubulin antibody, L. David Sibley for the MIC2, SAG1, and ALD antibody, Peter Bradley for the ROP1 antibody, Drew Etheridge for the GAP45 antibody, and Dominique Soldati-Favre for the GAP45 antibody. This work relied on VEuPathDB.org and we thank all contributors to this resource. This research was supported by funds from National Institutes of Health grants to SL (R01AI144369) and MB and MT (R01AI123457), a National Science Foundation Graduate Research Fellowship to AWC and ALH (174530). MT received funding from the Francis Crick Institute which receives its core funding from Cancer Research UK (CC2132), the UK Medical Research Council (CC2132), and the Wellcome Trust (CC2132). The Science Technology Proteomics Platform at the Francis Crick Institute received funding from Cancer Research UK (CC0199), the UK Medical Research Council (CC0199), and the Wellcome Trust (CC0199). MATERIALS AVAILABILITY STATEMENT Proteomics data is deposited on ProteomeXchange Consortium via the PRIDE partner repository. Accession numbers can be found in the published manuscript on eLife. The sequences of cloning vectors and primers generated for this study are listed in the table below. Custom analysis scripts in the R computing language are available upon request. Strains generated for this study are available upon request. 93 CHAPTER 4: Conclusions This thesis describes the identification and characterization of the HOOK complex as a regulator of microneme exocytosis in T. gondii. Chapter 2 describes two quantitative mass spectrometry approaches to identify substrates of the CDPK1 kinase. This resulted in the identification of 163 proteins phosphorylated in a CDPK1-dependent manner. A subset of substrates have previously characterized functions during motile stages, but many candidates remain uncharacterized. Chapter 3 establishes a partially conserved FHF complex that mediates rapid microneme trafficking in motile parasites. Loss of the complex results in defects in gliding and invasion. This is the first activating adaptor complex discovered in apicomplexans and represents a significant advance in understanding how micronemes may be linked to cortical microtubules and the regulation of their trafficking. Characterization of additional CDPK1 substrates will be required to fully understand how the kinase controls parasite motile stages. As there are many targets, systematic methods to screen or prioritize functionally relevant candidates will be required. As mentioned previously, the kinome of T. gondii has previously been tagged with a functionalized module enabling localization and conditional protein depletion(Smith et al. 2022). This method can be readily adapted to all CDPK1 targets to enable phenotypic screening in pooled and arrayed formats. Egress and invasion can be assessed to identify relevant targets involved in motile stages before more in-depth characterization that would include assessing microneme protein secretion and additional molecular mechanisms. Alternative biochemical approaches exist to prioritize candidates. Hotspot Thermal Profiling combines established methods for cellular thermal proteome profiling and enrichment methods for phosphorylated peptides(Huang et al. 2019). This quantitative proteomic method derives melting temperature for proteins subjected to different contexts, in which a shift in melting temperature suggests altered stability of a protein as a result of protein or ligand binding, for example. Cellular thermal shift assays have already been adapted to T. gondii to identify the Ca2+-responsive proteome in tachyzoites, so the addition of enrichment for phosphorylation is feasible (Herneisen et al. 2022). Bioinformatically, structure prediction algorithms such as 94 AlphaFold have provided predicted structures for proteins in T. gondii(Jumper et al. 2021). PTMs have been mapped onto predicted structures to predict potential functional consequences of these modifications in 3D space (Bludau et al. 2022). While AlphaFold structures do not dynamically change as a result of adding PTMs, mapping these modifications may help localize sites to identify the potential for conformational changes or protein-binding interfaces that regulate the activity. Characterizing the functional importance of PTMs at scale remains a major bottleneck in understanding signaling pathways. The development of systematic genetic and biochemical tools will significantly accelerate PTM characterization. I characterized two of the four members of the T. gondii HOOK complex. Similar characterization performed in this study will need to be applied to the uncharacterized binding partners. Neither of the two partners resemble the conserved FHIP, suggesting parasite-specific features. TGGT1_306920 is fitness conferring based on loss of function screens, whereas TGGT1_316650 appears dispensable. This suggests that TGGT1_306920 may serve a similar function as HOOK and FTS. TGGT1_316650 may indeed be dispensable, but the loss of function screens do not necessarily capture genes that may negatively regulate motile stages. One hypothesis is that TGGT1_316650 negatively regulates the HOOK complex to block microneme trafficking, thus loss of function mutations would not necessarily lead to fitness consequences. As many eukaryotic models utilize a conserved FHF complex, determining the unique features of this divergent complex may reveal new mechanisms for activating adaptors. Many aspects regarding the molecular mechanisms that enable rapid microneme trafficking during parasite motility still remain. Currently, the highest experimental standard for demonstrating a candidate adaptor activates dynein-dynactin motility is by reconstituting motility from purified components (Reck-Peterson et al. 2018). Immobilized microtubules are combined with fluorescently labeled dynein-dynactin, followed by the addition of a candidate adaptor. Total internal reflection microscopy is used to monitor activated motility along microtubules, enabling quantitative measurements of motor landing rates, velocity, and processivity. Cell-free reconstitution of adaptor-activated dynein-dynactin motility has been demonstrated for the human 95 adaptors BICD2, HOOK3, ninein and ninein-like (Reck-Peterson et al. 2018). In T. gondii, reconstituting adaptor-activated motility is currently not possible, as virtually all components of the cytoplasmic dynein-dynactin complex remain unidentified or uncharacterized. Eukaryotes typically express a single cytoplasmic dynein heavy chain. In T. gondii, TGGT1_294550 is the top candidate for the cytoplasmic dynein heavy chain (DHC) as it contains the necessary functional domains, is conserved among apicomplexans, and is required for parasite fitness based on genome-wide knockout screen data. Several other members of the dynein-dynactin complex have been predicted based on sequence homology(Gordon and Sibley 2005). The T. gondii genome also encodes other putative DHCs that are dispensable for the asexual stages of the parasite, but may be axonemal dyneins required for flagellar function in sexual stage parasites. Characterization of the dynein-dynactin complex within T. gondii will provide opportunities to identify additional candidate activating adaptors. Proximity labeling mass spectrometry of the dynein intermediate chain identified several activating adaptors in human cells and such methods can be applied to T. gondii to reveal adaptors for other vesicular cargo (Redwine et al. 2017). It also remains unknown what the HOOK complex binds to on micronemes to mediate trafficking during motile stages. In H. sapiens and D. melanogaster, yeast two-hybrid assays, IP-MS, and proximity labeling experiments have identified RAB5 GTPase as an interaction partner of FHIP, thus mediating the connection between the HOOK complex and endosomal cargo. In T. gondii, RAB5 has been implicated in the biogenesis of micronemes and may bind the uncharacterized members of the HOOK complex speculated to mediate cargo binding (Kremer et al. 2013). Together, these outline significant questions regarding the identity of the cytoplasmic dynein complex and the factor linking the HOOK complex for microtubules While kinase activity of CDPK1 is a central node in activating microneme exocytosis, characterization of downstream pathway components has been limited. In this thesis, I identified kinase-substrate interactions using quantitative proteomic approaches. This revealed connections with established regulators of motile stages, but also generated hypotheses implicating uncharacterized factors during parasite motility. 96 Kinase-mediated protein phosphorylation is a fundamental mechanism coordinating cellular processes. Identification of kinase targets can find new components of critical pathways. This was demonstrated in validating the HOOK complex as a new regulator of microneme exocytosis. Interestingly, parasites lacking HOOK do not mimic CDPK1 phenotypes exactly. Parasites depleted of HOOK are still able to initiate gliding and egress, but cannot sustain gliding nor invade new host cells. This suggests that HOOK functions comprise a subset of CDPK1-regulated activities. I hypothesize that a subpopulation of micronemes are docked within the apical complex and do not require HOOK-mediated trafficking. 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Abdelfattah, et al. 2011. “An Expanded Palette of Genetically Encoded Ca2+ Indicators.” Science 333 (6051): 1888–91. 117 SUPPLEMENTARY DATA Table 1 - Primers used in this work Primer publication ID Primer name Sequence AGCGAAGAAGATCTGTAACCCGGGCATATGTAGAAAAGTTGTAACGTTA P1 o14 GTAAACGTAAC P2 o15 AATCAGTTTCTGTTCAGAGCCGCCGCCGCCACT P3 o12 AACTTGACATCCCCATTTAC P4 o16 GAGAATAGACGTCACGCACAGTTTTAGAGCTAGAAATAGC CTTTGACGAGTTTCAACAGATGCTCTTGAAGCTCTGCGGAAATAGCAAG P5 o35 GGCTCGGGCTC GGGCTGTGAAGTCGGAGAGGGCCACCGCAGGATTTACGGCATAGGGCG P6 o36 AATTGGAGCTCC P7 o17 GTTGTCCCATTTTCTCGCGTTCGATAGAAAAGC P8 o18 CCGGACTACGCGTAGTTAATGCGGAGCTGGTAAGGAGC P9 o19 CTCCGGATCCAGCGTAATCTGGAACATCGTATG P10 o20 ACGCGAGAAAATGGGACAACAAGAGAGTAC P11 o21 ACTTTCGTCGTAGTCTTAATTTACTTGTACAGCTCGTC P12 o22 AGATTACGCTGGATCCGGAGAAGGAAGAG P13 o23 TCTCTTGTTGGGCCATTTTCTCGCGTTCGATAGAAAAGC P14 o24 CTCCTGATCCAGCGTAATCTGGAACATC P15 o25 ACGCGAGAAAATGGCCCAACAAGAGAGTACGTTG P16 o26 AGATTACGCTGGATCAGGAGAAGGAAGAG TCCGTTTTTTTGTCGTGTCGCGAAATCGTTCTTCTGCAAAATGTATCCCTA TGACGTGCCTGATTACGCCGGTGGCGGAGGCTCGgagaagagcgcgtgtccta aagatcccgctaagccgcctgccaaggcccaggtggttggctggcccccggttaggagttaccg caagaacgtgatggtatcttgccagaagtctagtggtggccctgaggcggcggcattcgttaaa gtctccatggacggagcgccgtacctgcgaaagattgatttgcgaatgtataaaagtTCTGG AAGTGAAACGCCTGGGACAAGCGAATCCGCAACTCCCGAGAGTATGTCT P17 gAWC012 TTCACTCACGCGCCCTTGCTCGACACAGATGCGT P18 AWC016 TCCGTTTTTTTGTCGTGTCG P19 AWC059 ACGCATCTGTGTCGAGCAAG TAAATGGGGATGTCAAGTTgGTCACGCGCCCTTGCTCGACAgttttagagcta P20 AWC061 gaaatag ctatttctagctctaaaacTGTCGAGCAAGGGCGCGTGACcAACTTGACATCCC P21 AWC062 CATTTA GTAAATGGGGATGTCAAGTTCGGGAGGCGTGAAAGGACTGGTTTaAGA P22 gAWC006 GCTAtgctgGAAAcagcaTAGCAAGTTtAAATAAGGCTAGTCCGTTATCAAC 118 TTGAAAAAGTGGCACCGAGTCGGTGCTTTTTTTTTCTTTTTCtctagaggtac CATGCATctaAGTCAGCGTCTCCAAAAGTAGAAGGGGCACTTCGAAGTCA AGAGACCGGTCTCACGACTCGATTGCGACAGATTCTGTCACACCTCGGG AGGCGTCTGGAAGTGAAACGCCTGG GTAAATGGGGATGTCAAGTTTCATTCGGCGTTGAAGAGGTGTTTaAGAG CTAtgctgGAAAcagcaTAGCAAGTTtAAATAAGGCTAGTCCGTTATCAACT TGAAAAAGTGGCACCGAGTCGGTGCTTTTTTTTTCTTTTTCtctagaggtacC ATGCATctaTGAGCTCAGCGAATAATTATCCGAAGCAGGTCACTCGGTAA GAGACCGGTCTCAGTCGGAAGGCAGTACTGGCGCGAACCTCTTCAACGC P23 gAWC011 CGAATCTGGAAGTGAAACGCCTGG P24 oALH426 tgcattcgctccggttttgcacagccgttctgtctcacgagaacgcatagtgttcgtccg P25 oALH091 tgtcggtgggtgccggctcgtctgtcgcaggtggcctgcagtgtcactgtagcctgccag CTCGTTCGGACCCCGCCCAAGCATGGGAGGCTGGCGCGGATCCGGACTC P26 AWC112 GGATCCGTGAGCAAGGGCGAGGAGGATAAC AACTTGGGCTCTCGTACGTGGAGAGACCTGTCTCCTTTCACTTGTACAGC P27 AWC113 TCGTCCATGCCCATCA TGGGGATGTCAAGTTGGGAGGCTGGCGCGGATGAAGTTTTAGAGCTAG P28 P102 AA P29 P103 TTCTAGCTCTAAAACTTCATCCGCGCCAGCCTCCCAACTTGACATCCCCA P30 o37 GAAACGGCGTGTAGCGTTTT P31 o38 TCTGGCAAGAGACCATCACG P32 o39 AACGAGATGTTCCGCGACTT P33 o40 GTGATTGACGCCGAGAGGAA P34 o41 GAAACGGCGTGTAGCGTTTT P35 o42 TCGGTCTCTTCGTGCACTC P36 o49 AGTGTGTGGACACGGATTCT P37 o50 CCGGACGTCAGTTGGTTGTAT P38 o51 AATACTTGACCAGCTGGCGT P39 o52 TATGGAAGCATGTGCGGCTC P40 o47 AGGACAAGGCAATTAACCATCTTG P41 o48 AGGACGTGAGACCGAACAGC P42 AWC058 GCACATTCGGTCCTCTCGAC P43 AWC021 gagaagatgccttgcaagtc P44 AWC014 GCGTTGACCCCAAAGAAGTT P45 oALH213 CGCTTGGAAGTACAGGTTTTC P46 oEC064 aaagagattttcgggattagccgg 119 P47 DS103 CAGAAACCGTCGCGTTTTCC P48 oSS318 ccctcgttcggaccccgcccaagcatg P49 oSS319 caacttgggctctcgtacgtggagagac 120 Table 2 - Plasmids used in this work GenBank ID/Addgene #/PubMed Plasmid Name ID Use pTUB-YFP-mAID- pTUB1_YFP_mAID_3HA Addgene: 87259 3xHA template template for iKD pTUB1_YFP_mAID_Myc GenBenk: Pending CDPK1 tagging Cas9 expression pSag1_Cas9-U6_sgUPRT Addgene: 54467 plasmid template for cWT CDPK1 pUPRT_CDPK1_ HA_T2A_GFP GenBank: Pending complementation templated for cMut CDPK1 pUPRT_CDPK1(G2A)_HA_T2A_mCherry GenBank: Pending complementation pUPRT_HA PMID: 21436047 pUPRT_HA empty Cas9- pSS013 GenBank: OM640003 expression plasmid Cas9-expression pSS014 GenBank: OM640002 plasmid V5-3xHA tagging pALH047 GenBank: ON312868 vector V5-mAID-HA tagging pALH086 GenBank: ON312869 vector gRNA/Cas9 plasmid targeting a neutral pBM041 GenBank: MN019116.1 locus template to amplify pMIC2-mNG-TurboID- pALH184 GenBank: Pending Ty-3'DHFR TurboID-Ty tagging pALH173 GenBank: Pending vector V5-mNG-mAID-Ty pGL015 GenBank: OM640005 tagging vector 121 Table 3 - Key resources table Reagent type (species) or Designatio Source or resource n reference Identifiers Additional information Strain, strain !"#$%!&#'()*+#'",-.! background (T. PMID: gondii) TIR1 28465425 $ doi: https://doi. !"#$%!&#'()*+#'",-.! Strain, strain org/10.110 $#/0.(&1231 background (T. CDPK1- 1/2022.07. gondii) AID 19.500742 45678-966814:%01$; !"#$%!&#'()*+#'",-.! Strain, strain $#/0.(&14:%01 background (T. TGGT1_3 gondii) iKD CDPK1 This paper 01440 <;=#",-.!$ !"#$%!&#'()*+#'",-.! $#/0.(&14:%01 Strain, strain <;=#",-.!$#/0.(&1":1 background (T. cWT TGGT1_3 gondii) CDPK1 This paper 01440 $>:1-?. !"#$%!&#'()*+#'",-.! $#/0.(&14:%01 Strain, strain <;=#",-.!$#/0.(&@->: background (T. cMut TGGT1_3 gondii) CDPK1 This paper 01440 A1":1$>:14/B699; Strain, strain 'CD*+'BEF background (T. gondii) G9HIJ?. !"#'()*+#'",-.!$#J?. Strain, strain background (T. PMID: gondii) CDPK1G 19218426 !"#'()*+#'",-.!$ Strain, strain !"#'()*+#'",-.!$#/0. background (T. PMID: TGGT1_3 gondii) CDPK1M 23149386 01440 (&@-&>*145678-96681 background (T. mNG- gondii) TurboID This paper $D9N7%01$; Strain, strain !"#'()*+#'",-.!$#/O: background (T. CLAMP- PMID: TGGT1_2 gondii) mNG 27594426 65790 <.145678-9668 Strain, strain !"#$%!&#'()*+#'",-.! background (T. TIR1/CLAM TGGT1_2 gondii) P-mNG This paper 65790 $#/O:<.145678-9668 !"#$%!&#'()*+#'",-.! TIR1/CLAM $#":14:%01 Strain, strain P- "KK(#/O:<.1 background (T. mNG/AID- TGGT1_2 gondii) HOOK This paper 65790 45678-9668 Human Foreskin Cell line (Homo Fibroblasts SCRC- sapiens) (HFFs) ATCC 1041 Mouse Provided by Matthew Child Antibody polyclonal other and Matt Bogyo. WB 123 anti- (1:3000). Only used in CDPK1 Figure 2. Mouse Cat# 05- monoclonal 724, clone 4A6 RRID:AB_ Antibody anti-Myc Millipore 11211891 WB (1:1000) Cat# Rat 11867423 monoclonal 001, clone 3F10 RRID:AB_ WB (1:1000); IFA (1:1000). Antibody anti-HA Roche 390919 Only used in Figure 2. Mouse monoclonal clone TP3 Cat# anti- ab8313, Toxoplasm RRID:AB_ Antibody a Abcam 306466 WB (1:1000) Rabbit Provided by Peter Bradley polyclonal Lab. WB (1:1000). Only Antibody anti-GAP45 other used in Figure 2. Cat# 11814460 Mouse 001, monoclonal RRID:AB_ Antibody anti-GFP Roche 390913 WB (1:1000) Rabbit Cat# polyclonal ab167453, anti- RRID:AB_ Antibody mCherry Abcam 2571870 WB (1:1000) Provided by John Rabbit Boothroyd Lab; WB monoclonal (1:10,000). Only used in Antibody anti-SAG1 other Figure 2. Rabbit monoclonal clone 51-8 Cat# anti- ab92570, thiophosph RRID:AB_ Antibody ate ester Abcam 10562142 WB (1:5000) 124 Developme ntal Mouse Studies monoclonal Hybridoma clone Bank at the RRID: 12G10 anti- University AB_11579 Provided by Marc-Jan Antibody tubulin of Iowa 11 Gubbels Lab. WB (1:2000) Guinea pig monoclonal anti- Custom Antibody CDPK1 Covance antibody WB (1/50,000) Rabbit Cat# monoclonal Cell 3724, (C29F4) Signaling RRID:AB_ Antibody anti-HA Technology 1549585 WB (1:1000); IFA (1:1600) Mouse monoclonal Provided by L. David clone 6D10 PMID: Sibley Lab. WB (1:5000); Antibody anti-MIC2 10799515 IFA (1:2000) Mouse Provied by Peter Bradley Antibody anti-ROP1 other Lab. IFA (1:2000) Mouse Provided by L. David polyclonal PMID: Sibley Lab. IFA (1:500). Antibody anti-SAG1 3183382 Used for invasion assays. Provided by R. Drew Rabbit Lampire Etheridge Lab. IFA polyclonal Biological (1:1000). Used for invasion Antibody anti-GAP45 Laboratory assays. Cat# Mouse 901533, monoclonal RRID: (16B12) AB_25650 Antibody anti-HA BioLegend 05 WB (1:1000) Rabbit polyclonal clone WU1614 PMID: Provided by L. David Antibody anti-ALD 16923803 Sibley Lab. WB (1:10,000) Rabbit Provided by Dominique polyclonal Soldati-Favre Lab. WB Antibody anti-GAP45 other (1:5000); IFA (1:5000) 125 Peroxidase -AffiniPure Jackson polyclonal ImmunoRe Cat# 111- Goat Anti- search 035-003, Rabbit IgG Laboratorie RRID:AB_ Antibody (H+L) s 2313567 WB (1/5000) Alexa Fluor Life 594 anti- Technologi Antibody rabbit es IFA (1:1000) Alexa Fluor Life 488 anti- Technologi Antibody mouse es IFA (1:1000) Alexa Fluor Life 488 anti- Technologi Antibody rabbit es IFA (1:1000) Alexa Fluor Life 594 anti- Technologi Antibody mouse es IFA (1:1000) IRDye 800CW Donkey anti-Guinea Pig IgG Secondary LICOR:92 Antibody Antibody LICOR 6-32411 WB (1:10,000) IRDye 680RD Donkey anti-Guinea Pig IgG Secondary LICOR:92 Antibody Antibody LICOR 6-68077 WB (1:10,000) IRDye 800CW Goat anti- Mouse IgG1- Specific Secondary LICOR:92 Antibody Antibody LICOR 6-32350 WB (1:10,000) IRDye LICOR:92 Antibody 680LT Goat LICOR 6-68020 WB (1:10,000) 126 anti-Mouse IgG Secondary Antibody IRDye 800CW Goat anti- Rabbit IgG Secondary LICOR:92 Antibody Antibody LICOR 6-32211 WB (1:10,000) IRDye 680LT Goat anti-Rabbit IgG Secondary LICOR:92 Antibody Antibody LICOR 6-68021 WB (1:10,000) Alpaca clone 1B7 anti- PMID: Antibody CDPK1 26305940 n/a IRDye 680RD Peptide, recombinant Streptavidi LICOR:92 protein n LICOR 6-68079 WB (1:3,000) Peptide, recombinant PMID: protein Aerolysin 26584919 n/a Sequencin g Grade Peptide, recombinant Modified Promega: protein Trypsin Promega V5113 3- Indoleaceti Sigma Chemical compound, c acid Sigma Aldrich:I28 drug (auxin) Aldrich 86-5G Chemical compound, Compound PMID: drug 1 12455981 n/a Santa Chemical compound, Hoechst Cruz:sc- drug 33258 Santa Cruz 394039 Egress assay (1:4000) Chemical compound, Hoechst Invitrogen: drug 33342 Invitrogen H3570 IFA (1:20,000) 127 DAPI (4',6- Diamidino- 2- Phenylindol e, Chemical compound, Dihydrochl Invitrogen: drug oride) Invitrogen D1306 Thermo Chemical compound, Prolong Thermo Fisher:P3696 drug Diamond Fisher 5 Chemical compound, Calbioche Calbiochem:6 drug zaprinast m 84500 Chemical compound, Calbioche Calbiochem:1 drug A23187 m 00105 Tokyo Chemical compound, Myristic Chemical Cat# drug acid Industry M0476 Alkyne- myristic Chemical compound, acid Cat# RL- drug (YnMyr) Iris Biotech 2055 Trypsin PMID: cleavable 25807930; This reagent was first Chemical compound, capture PMID: reported as RTB in PMID: drug reagent 32618271 25807930. Thermo Thermo Fisher Chemical compound, L-Proline Fisher Scientific: drug for SILAC Scientific 88211 Fetal Bovine Serum, dialyzed, US Thermo origin, One Thermo Fisher Chemical compound, Shot™ Fisher Scientific: drug format Scientific A3382001 Thermo Thermo Fisher Chemical compound, DMEM for Fisher Scientific: drug SILAC Scientific 88364 L-Arginine- Thermo Chemical compound, HCl for Fisher Thermo drug SILAC Scientific Fisher 128 Scientific: 89989 Thermo L-Lysine- Thermo Fisher Chemical compound, 2HCl for Fisher Scientific: drug SILAC Scientific 89987 L-Arginine- Thermo HCl, 13C6, Thermo Fisher Chemical compound, 15N4 for Fisher Scientific: drug SILAC Scientific 89990 L-Lysine- 2HCl, Thermo 13C6, Thermo Fisher Chemical compound, 15N2 for Fisher Scientific: drug SILAC Scientific 88209 p- nitrobenzyl Chemical compound, mesylate Epitomics drug (PNBM) Epitomics # 3700-1 OXONE®, Sigma monopersu Aldrich: Chemical compound, lfate 228036- drug compound Sigma Aldrich 5G N6- furfurylade nosine (kinetin)-5′- O-[3- thiotriphos phate] Axxora:BL Chemical compound, sodium salt G-F008- drug (KTPγS) Axxora 05 Adenosine 5′- triphosphat e disodium Sigma Chemical compound, salt hydrate Sigma Aldrich:A6 drug (ATP) Aldrich 419-1G Guanosine 5ʹ- Chemical compound, Triphospha Calbioche Millipore:3 drug te, m 71701 129 Disodium Salt (GTP) β-Casein Sigma Chemical compound, from Sigma Aldrich:C6 drug bovine milk Aldrich 905 Sigma Chemical compound, Sigma Aldrich: drug Biotin Aldrich B4501-1G PP1 Analog Sigma Chemical compound, III, 3-MB- Calbioche Aldrich:52 drug PP1 m 9582 Protific:C0 Commercial assay or S-trap 2-micro- kit micro Protifi 80 Pierce Quantitativ e Thermo Fluorometri Thermo Fisher Commercial assay or c Peptide Fisher Scientific: kit Assay Scientific 23290 Thermo TMTpro™ Thermo Fisher Commercial assay or 16plex Label Fisher Scientific: kit Reagent Set Scientific A44522 EasyPep™ Thermo MS Sample Thermo Fisher Commercial assay or Prep Kits - Fisher Scientific: kit Maxi Scientific A45734 High-Select TiO2 Phosphope Thermo ptide Thermo Fisher Commercial assay or Enrichment Fisher Scientific: kit Kit Scientific A32993 High- Select™ Fe- NTA Phosphopep Thermo tide Thermo Fisher Commercial assay or Enrichment Fisher Scientific: kit Kit Scientific A32992 130 Pierce High pH Reversed- Phase Thermo Peptide Thermo Fisher Commercial assay or Fractionati Fisher Scientific: kit on Kit Scientific 84868 Dynabeads® MyOne™ Commercial assay or Streptavidin Cat# kit C1 Invitrogen 65001 Bio- Commercial assay or Bio-Rad Rad:5000 kit DC assay Bio-Rad 116 Sep-Pak C18 Plus Short Cartridge, 360 mg Sorbent per Commercial assay or Cartridge, Waters:W kit 55-105 µm Waters AT020515 Thermo SulfoLink™ Thermo Fisher Commercial assay or Coupling Fisher Scientific: kit Resin Scientific 20401 Radiance Plus Chemilumi Commercial assay or nescent Azure VWR:1014 kit Substrate Biosystems 7-298 Pierce™ Anti-HA Thermo Commercial assay or Magnetic Thermo Scientific: kit Beads Scientific 88836 Pierce™ Streptavidin Thermo Commercial assay or Magnetic Thermo Scientific: kit Beads Scientific 88817 131 All plasmids used in this study are listed in Recombinant DNA Supplemen reagent tary file 10 All primers and oligonucleo tides used in this study are listed in Sequence-based Supplemen reagent tary file 10 Proteome Discoverer Thermo Software, algorithm 4.2 Fisher MaxQuant (version Free software for searching 1.5.0.25 PMID: RRID:SCR of mass spectrometry Software, algorithm and 1.5.2.8) 19029910 _014485 acquisition files Perseus Free software for (version PMID: RRID:SCR processing of MaxQuant Software, algorithm 1.5.0.9) 27348712 _015753 output files Scaffold Proteome Software, algorithm DIA Software R Foundation for R version Statistical Software, algorithm 4.0 Computing Software, algorithm Prism 8 GraphPad PMID: Software, algorithm Fiji 22743772 PMID: Software, algorithm ilastik 31570887 ilastik.org PMID: Software, algorithm HHPRED 29258817 132 https://ww w.snapge Software, algorithm SnapGene other ne.com/ PMID: ToxoDB.o Software, algorithm ToxoDB 18003657 rg DMEM, Life high Life Technolog glucose Technologi ies:11965 other (media) es 118 HALT Thermo protease Thermo Fisher:877 Materials and Methods: other inhibitor Fisher 86 lysis buffer HALT protease and Thermo phosphatas Thermo Fisher:PI7 Materials and Methods: other e inhibitor Fisher 8440 lysis buffer Sigma Sigma Aldrich:E1 Materials and Methods: other Benzonase Aldrich 014 lysis buffer 133