Deciphering the Substrate Selectivity of a Rhomboid

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A dissertation submitted to the Johns Hopkins University in conformity with the requirements of the degree of Doctor of Philosophy

Baltimore, MD March, 2018 $EVWUDFW

The rhomboid family of intramembrane serine proteases has been implicated in critical parasite functions such as immune evasion, host-cell attachment, and, in the case of

ROM4 from Plasmodium falciparum, invasion. Because this process is unique to parasites,

PfROM4 represents an attractive therapeutic target. Targeting, however, is complicated by the fact that PfROM4 exhibits atypical selectivity: PfROM4 cleaves parasite but not canonical rhomboid substrates. Moreover, invasion is rapid (<1 minute), and it remains unclear how ‘druggable’ the membrane-immersed rhomboid family of proteases are.

I mapped the basis of PfROM4’s atypical selectivity to the juxtamembrane sequence of substrates and found that selectivity operates by steric exclusion – PfROM4 can cleave a diversity of sequences derived from parasites but cannot cleave canonical substrates due to specific steric clashes between substrate and enzyme. This represents a distinct mechanism for achieving substrate specificity in a membrane-immersed environment.

Exploiting the rules governing PfROM4’s substrate selectivity, I was able to generate

‘super-substrate’ mutants that enhanced proteolysis ~5-20 fold. Our laboratory also identified boronate as a warhead that enhanced inhibition ~100-fold. I designed a ‘super- substrate’ derived peptide-boronate compound and found that it selectively inhibited

PfROM4 proteolysis with μM potency.

Added to malaria cultures, this compound disrupted host-cell invasion, and parasite growth ceased over 2-3 cycles. Not only does this work establish for the first time that

PfROM4’s activity is necessary and targetable for parasite invasion, it also provides a strategy for designing selective rhomboid inhibitors for enzymes from other organisms.

Thesis Advisor: Sinisa Urban, PhD

Thesis Reader: Sean Prigge, PhD

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To the individuals along this journey that I have been fortunate to call teachers and mentors: My high school biology teacher, Bob Braddy, piqued my interest in research and taught me that, at its root, science is an endeavor in curiosity. Through their example, former mentors Dr. Robert Siliciano and Dr. Joel Blankson, not only taught me to pursue important questions, but to do so with dedication and humility.

The Medical Scientist Training Program (MSTP) and Biochemistry, Cellular and

Molecular Biology (BCMB) Gradate Program have provided the flexibility and support that enabled me to complete this joint-degree program. I am thankful to the faculty, students, and administrators in these programs for creating an environment that allowed me to grow both professionally and personally.

I am also thankful to members of my Thesis Committee and the Molecular Biology and Genetics department for providing feedback that strengthened this story and helped to make me a better scientist.

I am grateful to both current and former members of the Urban Lab for their insight, help, and friendship. I am particularly indebted to my thesis mentor, Dr. Sinisa

Urban. Through his enthusiasm for his science, fearlessness in tackling difficult questions, and dedication to his family, he has set an example for what I aspire to be, both as a scientist and as a husband and father.

Ultimately, this work would not be possible without my wife Renuka, parents, or brother Pujan. They set the stage for all that I have accomplished by encouraging my curiosity and providing unconditional support throughout this long journey. I am proudest to be a husband to Renuka and a father to Aarav, and I am excited to see where this journey of life takes us.

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Abstract ii

Acknowledgments iii

Table of Contents iv

List of Tables v

List of Figures v

Chapter I: Introduction 1

Protozoan Parasites 2

Apicomplexan Parasites 3

Invasion in Apicomplexa 4

Adhesins are ‘shed’ from the merozoite surface 8

Rhomboid proteases are the intramembrane ‘sheddases’ 9

A chemical biology approach to target PfROM4 11

References 14

Figure Legends 31

Chapter II: A steric exclusion mechanism governs rhomboid substrate

selectivity 33

Summary 34

Introduction 35

Results 40

Discussion 51

Materials and Methods 53

Acknowledgements 55

References 55

Figure Legends 59

Chapter III: PfROM4 is druggable and essential for invasion by blood-stage parasites 75

Summary 76

Introduction 77

Results 77

Discussion 85

Acknowledgments 85

Materials and Methods 86

References 91

Figures Legends 96

Curriculum Vitale 109

List of Tables

Table 2.1 71

Table 2.2 72

Table 2.3 73

Table 2.4 73

Table 2.5 73

Table 2.6 74

Table 2.7 74

List of Figures

Figure 1.1 32

Figure 2.1 64

Figure 2.2 65

Figure 2.3 66

Figure 2.4 67

Figure 2.5 68

Figure 2.6 69

Figure 2.7 70

Figure 3.1 101

Figure 3.2 102

Figure 3.3 103

Figure 3.4 104

Figure 3.5 105

Figure 3.6 106

Figure 3.7 107

Figure 3.8 108

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Protozoan parasites

Protozoan parasites are an extraordinarily diverse group of ancient human and animal pathogens that occupy many phyla in the eukaryotic tree of life. Protozoa, meaning

‘first animals’, were historically grouped into four polyphyletic groups based on their morphology and means of locomotion: the ameobae, flagellates, cilates and sporozoa

(Honigberg et al. 1964). Given that the strategy of parasitism has evolved independently multiple times within these groups, protozoan parasites share little aside from a nucleus and the enormous burden they impart on society (Poulin and Randhawa 2015): the amoebic parasites of the Entamoeba family are a global cause of diarrheal disease (Haque et al. 2003); trypanosomal flagellates are the cause of both African sleeping sickness (Brun et al. 2010) and Chagas disease (Rassi, Rassi, and Marcondes de Rezende 2012); there are >2,500 ciliated parasites that cause mostly animal disease (Lynn 2010); finally, the group formally known as sporozoa (now apicomplexa) includes many important human pathogens such as Plasmodium, the causative agent of malaria. Collectively, these protozoa exact a human health burden greater than that of HIV/AIDS (Hay et al. 2017).

Despite this societal importance, our understanding of molecular mechanisms of pathogenesis for most protozoan parasites remains limited. The reasons are both economic and technical: First, parasitic diseases disproportionately affect disadvantaged populations and have consequently suffered from a lack of research funding and attention (Hotez et al.

2007). Secondly, these parasites are among the most complicated single-cell organisms in existence - separated by hundreds of millions of years of evolution from opisthokonts

(animals, yeast), they contain structures for metabolism, locomotion, and invasion that are unique in the eukaryotic domain (Baldauf 2003). Finally, many of the laboratory tools developed to study higher eukaryotes have not been optimized to study these organisms.

Apicomplexan parasites

Of all protozoan parasites, those from the phylum Apicomplexa represent the greatest threat to human health. One of its members, Plasmodium spp., is the causative agent of malaria, a disease that is still responsible for ~700,000 deaths per year and ~60 million disability adjusted life years (DALYs) (Naghavi et al. 2017; Hay et al. 2017). Other medically relevant members include Toxoplasma gondii, which causes neurological diseases in fetuses and immunocompromised adults (Wohlfert, Blader, and Wilson 2017), and Cryptosporidium parvum, a leading cause of childhood diarrhea (Checkley et al. 2015). Another member,

Babesia, is an emerging cause of hemolytic anemia in the NE United States and Europe

(Vannier et al. 2015).

Yet, therapies for most diseases caused by apicomplexan parasites remain lacking: there is an absence of effective therapies for Cryptosporidiosis altogether (Smith and

Corcoran 2004); that there is only a single anti-toxoplasmosis agent was illuminated by the recent scandal over Turing Pharmaceutical’s exorbitant price increase for pyrimethamine

(Gallant 2015), an action that would be impossible had there been other effective therapies on the market; and, in malaria, recent progress has been threatened by spreading resistance to the first-line therapy artemisinin (Packard 2014; Dondorp et al. 2009; Yeung et al. 2009;

Ashley et al. 2014; Noedl et al. 2008; Wells, Hooft van Huijsduijnen, and Van Voorhis 2015).

Moreover, most of the drugs we do have were identified in phenotypic assays and their mechanisms of action, if known, were only discovered later (Wells, Hooft van

Huijsduijnen, and Van Voorhis 2015). This is in contrast to other infectious diseases like

HIV and HCV, where a thorough understanding of their biology led to the design of

3 mechanism-based inhibitors which have revolutionized treatment and saved the lives of millions of people (Walensky et al. 2006; Chung and Baumert 2014). Thus, determining the molecular mechanisms of pathogenesis for protozoan pathogens, and apicomplexan parasites specifically, would prove useful in the rational design of novel anti-parasitic chemotherapy.

Invasion in apicomplexa

Apicomplexan parasites have a complex lifecycle, shuffling between different hosts as a part of their development (Levine 1988). Though parasites take many different forms during this process, they are unified in their ability to invade other cells. Indeed, the phylum’s namesake is the apical complex, a group of secretory organelles tethered to a locomotive apparatus that are responsible for adhesion to and invasion of host cells

(Blackman and Bannister 2001). This ability to invade by forcing the invagination of the host cell membrane around the parasite is a feature that distinguishes members of Apicomplexa from other eukaryotes (L D Sibley 2004). It also accounts for the pathology that these parasites exhibit: Cryptosporidium invades enterocytes of the small intestine, triggering gastrointestinal symptoms (Striepen 2013). Toxoplasma causes neurological symptoms on account of its ability to invade almost any nucleated cell, most notably neurons (Dubey

2008). The synchronous egress and entry of erythrocytes by parasites from the Plasmodium spp. results in malaria’s hallmark periodic fever, and accounts for its morbidity and mortality

(Cowman and Crabb 2006).

While invasion is best understood in Toxoplasma gondii, where tractable genetics and ease of laboratory cultivation have allowed for the basic molecular details of this process to be identified (Kim and Weiss 2004; V. Carruthers and Boothroyd 2007), its general principles apply to other members of the phylum (L. M. Weiss and Kim 2013). Because

4 malaria is of great societal importance, and is the emphasis of my thesis, this outline focuses on how invasion (recently reviewed in (Cowman et al. 2017)) is thought to occur in

Plasmodium falciparum, the deadliest of the four Plasmodium species that routinely infect humans (Murray et al. 2012).

In the asexual, blood stage infection, the subsequent round of invasion begins as soon as merozoites egress from a rupturing, parasitized erythrocyte (Abkarian et al. 2011;

Glushakova et al. 2005; lifecycle depicted in Figure 1.1A). This invasive form of Plasmodium is uniquely suited to recognize, bind to, and invade new erythrocytes by forcing the invagination of the host cell membrane around it, forming the parasitophorous vacuole (PV).

It is here where the parasites grow, divide, and prepare for the next round of invasion

(Grüring et al. 2011). As they undergo maturation, though, the intraerythrocytic forms of P. falciparum express a number of endothelial cell binding molecules on the erythrocyte plasma membrane in order to sequester the parasitized erythrocyte in the microvasculature, thereby protecting themselves from splenic clearance (Miller et al. 2002). In this process, however, they also occlude blood flow to many end-organs and trigger deleterious immune responses, resulting in dreaded complications such as cerebral malaria (Sheehy and Reba 1967).

Thus, because of its unique nature in the eukaryotic domain, and its relation to pathology, erythrocytic invasion has been the subject of much investigation with the goal of disrupting this process as a therapeutic strategy (Miller et al. 2013). Targeting invasion rather than other essential pathways may allow for the development of specific inhibitors with few off-target effects as the invasion machinery is not present in host organisms. Yet, despite this promise, few invasion-specific inhibitors have been identified (Chandramohanadas et al.

2014; Pino et al. 2017), and there is a need to validate new therapeutic targets in this pathway

(Deu 2017).

Invasion itself is dependent on a series of intricate, highly orchestrated ligand binding events to the host-cell surface (partly dissected in Riglar et al. 2011; depicted in Figure 1.1B).

Initial binding to the host-cells is mediated by interactions between the abundant GPI- anchored parasite surface protein MSP1 and unidentified receptors on the host-cell membrane (Dzierszinski et al. 2000; Mineo and Kasper 1994; Goel et al. 2003; O’Donnell et al. 2000). This binding is characterized by low-affinity, reversible interactions. Following successful attachment, the parasite re-orients itself so that its apical end, which contains the invasion apparatus, is facing perpendicular to the plane of the host-cell membrane (Aikawa et al. 1978; Dvorak et al. 1975). At this point, two groups of intracellular organelles, micronemes and rhoptries, fuse to the apical end, releasing a number of type I transmembrane adhesins onto the parasite cell surface that mediate subsequent steps in invasion (Riglar et al. 2011; S. Singh et al. 2010; Richard et al. 2009).

In Plasmodium spp., two of these adhesin families, the erythrocyte-binding-like proteins (EBL) and the reticulocyte-binding protein homologues (Rh), commit the parasite to invasion by forming high-affinity interactions with their cognate erythrocyte receptors (S.

Singh et al. 2010; Riglar et al. 2011; Lopaticki et al. 2011; G. E. Weiss et al. 2015). The best characterized of these are EBA-175 and Rh4 which bind host proteins Glycophorin A and

Complement receptor 1, respectively (Sim et al. 1994; W.-H. Tham et al. 2010). They play a critical role in signaling parasites to release remaining secretory organelles onto the cell- surface, irreversibly committing the parasites to invasion. These families of adhesins mediate invasion through alternative pathways, serving both to counter host receptor polymorphism and to mediate evasion from the immune system (W. H. Tham, Healer, and Cowman 2012).

Thus, while EBA-175 and Rh4 are individually dispensable for invasion, they are collectively

6 essential, demonstrating that they play a redundant, but critical role in invasion (Duraisingh,

Triglia, et al. 2003; Stubbs et al. 2005; W.-H. Tham et al. 2010; Lopaticki et al. 2011).

Binding of the EBL/Rh proteins to their host-cell receptors signals the merozoite to begin its forced entry into the erythrocyte. Remarkably, the parasite translocates its own invasion aparatus onto the erythrocyte membrane in the form of the RON complex. One member, RON2, serves as an anchor for the parasite adhesin apical membrane antigen 1

(AMA1) to latch onto (Riglar et al. 2011; Cao et al. 2009; Collins et al. 2009; Richard et al.

2010; Tonkin et al. 2011; Olivieri et al. 2011; Lamarque et al. 2011; Mital et al. 2005; Triglia et al. 2000). The AMA1/RON interaction forms an electron dense complex between the parasite and erythrocyte plasma membranes, termed the ‘tight junction’, that serves as a molecular sieve, keeping erythrocyte membrane proteins out of the nascent parasitophorous vacuole (Riglar et al. 2011; Aikawa et al. 1978). Importantly, merozoite surface proteins, such as MSP1 and the EBL/Rh adhesins are also sequestered from the PV in this manner. The tight junction is connected to the parasite’s actin-mysoin motor (Baum et al. 2006;

Giovannini et al. 2011), which provides the propulsive force for the parasite to enter the erythrocyte.

Given their importance and abundance on the merozoite surface, much effort has been put into developing vaccines to these adhesins (reviewed in (Crompton, Pierce, and

Miller 2010)). Despite this push, efforts to-date have mostly failed for a number of reasons:

1) Adhesins mediate invasion through independent pathways - disrupting only one adhesin/receptor interaction is not sufficient to block invasion (A. P. Singh et al. 2005;

Duraisingh, Maier, et al. 2003). 2) Adhesins are highly polymorphic, and antibodies do not always cross-react with other haplotypes (Scherf, Lopez-Rubio, and Riviere 2008). 3) Even within an adhesin family, merozoites express many functionally redundant members on their

7 surface (W. H. Tham, Healer, and Cowman 2012). For these reasons, inducing an antibody response to a single adhesin is unlikely to be protective. Thus, alternate invasion-targeting strategies are needed.

Adhesins are ‘shed’ from the merozoite surface

An interesting observation is that the N-terminal ectodomains from the aforementioned adhesins (EBL, Rh, AMA1) can be found in the culture media of invading parasites (reviewed in Blackman 2004; Figure 1.1B). In fact, the EBL family was originally identified by their shed, soluble domains which bound Duffy determinates on erythrocytes

(Haynes et al. 1988). The ‘shedding’ of cell surface adhesins is mediated by proteolysis as it is sensitive to treatment by non-specific protease inhibitors (V. B. Carruthers 2006). Moreover,

‘shedding’ is thought to be essential both because treatment with protease inhibitors reduces invasion efficiency, and uncleavable mutants of the adhesins are often unable to be generated (Ejigiri et al. 2012; O’Donnell et al. 2006; Parussini et al. 2012; Child et al. 2010).

Proteolysis of these adhesins is thought to play two non-mutually exclusive roles: 1) To remove their bulky ectodomains from the parasite cell surface – termed ‘capping’ - thereby allowing the parasitophorous vacuole to seal at the posterior end (Figure 1.1B) and 2) To mediate immune evasion as the shed ectodomains can serve as decoys for host antibodies.

Over the past decade, much effort has been spent trying to characterize the proteases that mediate shedding, termed ‘sheddases’, with the goal of both understanding the role of proteolysis during invasion and targeting this process therapeutically. What makes ‘shedding’ particularly attractive from a therapeutic perspective is that this process appears to be universal to all invasion-mediating adhesins (Blackman 2004). Given how extensively the merozoite is coated with these adhesins (Bannister et al. 1975), blocking their release from

8 the parasite cell-surface could serve as an invasion-blocking strategy (Blackman 2004; Baker,

Wijetilaka, and Urban 2006).

While a subtilisin-like protease, PfSUB2, has been implicated in processing both

AMA1 and MSP1 (Barale and Blisnick 1999; Harris et al. 2005; S. A. Howell et al. 2003;

Figure 1.1C), extensive efforts to chemically disrupt its function have not succeeded (Deu

2017). Moreover, even the earliest studies of the effects of protease inhibitors on invasion suggested that there was more than one active protease in invasion (Hadley, Aikawa, and

Miller 1983). Thus, attention shifted to uncovering this second, undefined, ‘sheddase.’

Rhomboid proteases are the intramembrane ‘sheddases’

A surprising, but - at the time - controversial finding was that the cleavage site of this second sheddase mapped to the luminal side of transmembrane segments from Toxoplasma adhesins (Opitz et al. 2002). Contemporaneously, rhomboid proteins were characterized as intramembrane serine proteases that were uniquely capable of this unusual type of cleavage

(S Urban, Lee, and Freeman 2001). Rhomboid’s active site lies below the plane of the membrane, cleaving its substrates near the top of their transmembrane segments, thereby releasing the respective ectodomains from the cell surface (reviewed in (S Urban 2010)). Sin

Urban subsequently found that these Toxoplasma adhesins were substrates for non-parasitic rhomboid proteases and proposed that this unidentified ‘sheddase’ was, in fact, a yet-to-be discovered parasitic rhomboid protease (Sinisa Urban and Freeman 2003).

An ensuing study confirmed that the Toxoplasma adhesins were indeed cleaved within their transmembrane segments (X. W. Zhou et al. 2004). This unusual type of cleavage was also found to occur in the important Plasmodium adhesins EBA-175 and AMA1 (S. a Howell et al. 2005; O’Donnell et al. 2006), though in the case of AMA1, PfSUB2 cleavage was found to be the predominant mode of processing (S. a Howell et al. 2005). Finally, phylogenetic

9 evidence indicated that rhomboid may have unique functions in these parasites: While they are found in all domains of life (Koonin et al. 2003), rhomboid genes were found to have diversified in eukaryotic parasites such as those from Apicomplexa where the genomes of P. falciparum and T. gondii encode for 8 and 6 rhomboid-like genes, respectively (Dowse and

Soldati 2005; L. David Sibley 2013). Thus, a growing body of circumstantial data implicated rhomboid in the shedding of parasite adhesins.

Several lines of evidence subsequently linked rhomboid to the intramembrane proteolysis of adhesins during many parts of the parasite’s lifecycle, including invasion. First, while many rhomboid proteins from T. gondii were found to have enzymatic activity (Dowse et al. 2005; Brossier et al. 2005), one member, TgROM5, stood out as it was localized to the posterior end of the parasite cell surface and active against T. gondii adhesins in a heterologous cleavage assay (Brossier et al. 2005). Second, PfROM4, a TgROM5 ortholog from P. falciparum, cleaved a number of invasion-mediating adhesins from the EBL and Rh families, including EBA-175 and Rh4, in a similar heterologous cleavage assay (Baker,

Wijetilaka, and Urban 2006). Lastly, PfROM4 was found to be localized to the merozoite cell surface and proved refractory to attempts at genetic deletion (O’Donnell and Blackman

2005). Importantly, O’Donnell, et al. were also unable to generate parasites harboring a rhomboid-uncleavable version of EBA-175. For these reasons, TgROM5 and PfROM4 were identified as the putative intramembrane ‘sheddase’ in Toxoplasma and Plasmodium, respectively.

Despite over ten years of ensuing investigation, evidence linking PfROM4 to invasion remains largely indirect. While studies found roles for another Plasmodium rhomboid, ROM1, in liver stage disease (Srinivasan, Coppens, and Jacobs-Lorena 2009) and formation of the parasitophorous vacuole (Vera et al. 2011), ROM4’s precise function is

10 unelucidated as it remains refractory to deletion in blood-stage parasites (Lin et al. 2013).

Moreover, recent genetic evidence from Toxoplasma gondii has challenged the premise that

‘shedding’ is essential for invasion as a number of ROM knockouts, including TgROM5, proved dispensable for host-cell invasion (Shen et al. 2014; Rugarabamu et al. 2015). Though the respective adhesins accumulated on the cell surface, confirming that ROMs serve as

‘sheddases’, these ROM knockout parasites still invaded host-cells efficiently. Extending those findings, many in the field question whether PfROM4 is truly indispensable for invasion, attributing the accumulated indirect genetic evidence to technical issues.

A chemical biology approach to target PfROM4

Selective chemical inhibition of PfROM4 would provide a strategy to circumvent the inherent limitations of genetic approaches (Child 2013). Such inhibition could shed light on

PfROM4’s function in a time-resolved manner. Moreover, it would not only address

PfROM4’s essentiality in invasion, it could also answer a fundamental question in the field: are rhomboid enzymes therapeutically targetable (Deu 2017)?

Owing to their unique place in nature, rhomboid enzymes have proved difficult to inhibit chemically. Indeed, even in the first characterizations of rhomboids as intramembrane proteases, they proved refractory to almost all protease inhibitors save those harboring unstable masked electrophiles (S Urban, Lee, and Freeman 2001; Sinisa Urban and Wolfe

2005). Ensuing studies have highlighted many of the challenges in targeting these unusual enzymes (Figure 1.1C):

First, crystal structures of bacterial rhomboid enzymes in detergent revealed that while the serine-histidine catalytic dyad is water accessible and able to perform hydrolysis, rhomboid’s active site would nonetheless be buried when in native lipid bilayers (Wu et al.

2006; Wang, Zhang, and Ha 2006; Ben-Shem, Fass, and Bibi 2007). Though experimental

11 data in detergent indicated that the active site of bacterial rhomboid proteases may be accessible to small molecule probes (Maegawa et al. 2007), molecular dynamics simulations in lipid bilayers indicated that crystallographic waters are quickly lost (Y. Zhou et al. 2012), making it doubtful that the active site of rhomboid enzymes are easily accessible when in their native (lipid bilayer) environment. Moreover, eukaryotic enzymes such as PfROM4 harbor features outside of the 6-TM catalytic core that may form an additional barrier to entry. Specifically, PfROM4 has an unusually large Loop 1 segment that may overlay the enzyme’s catalytic core. Thus, a chemical inhibitor would have to navigate these structural constraints.

Second, rhomboid enzymes do not have physiological affinity for their substrates

(Moin and Urban 2012; Dickey et al. 2013). Rather, the dynamics of a substrate’s transmembrane segment governs rhomboid’s ability to cleave it (Moin and Urban 2012).

Indeed, co-crystal structures of rhomboid enzymes with substrate mimics have demonstrated few interactions between substrate sidechains and the enzyme (Zoll et al. 2014; Cho, Dickey, and Urban 2016). Given that the selectivity and potency of many protease inhibitors are derived from such interactions, rhomboid could prove challenging to inhibit in physiological settings.

Third, as discussed extensively in Chapter II, it is unclear how rhomboid enzymes achieve selectivity for their substrates. Although some have posited that rhomboid works like soluble proteases and recognize their substrates through targeting motifs (Strisovsky,

Sharpe, and Freeman 2009), our laboratory has found that many rhomboid enzymes appear to be sequence non-selective (Moin and Urban 2012). Without a mechanism to achieve substrate selectivity, it is unclear how to design a selective inhibitor. And given that there are emerging roles for rhomboid proteases in mammalian biology (Bergbold and Lemberg

2013), selective targeting of the parasitic rhomboid enzymes is a requirement. Here, though, there is hope as PfROM4 exhibits ‘atypical specificity,’ cleaving adhesins derived from parasites but not canonical substrates derived from bacteria or metazoans (Baker, Wijetilaka, and Urban 2006). Deciphering the mechanisms governing this ‘atypical’ recognition, then, could lead to the design of a parasite selective inhibitor.

Lastly, there is a narrow window of opportunity to inhibit PfROM4 as free merozoites rapidly invade (<1 minute) erythrocytes (Dvorak et al. 1975; Riglar et al. 2011).

Thus, the inhibitor must have rapid-on kinetics if it is to have a chance at inhibiting

PfROM4.

In recent years, there has been a number of attempts to design chemical inhibitors of rhomboid (reviewed (Wolf and Verhelst 2016)). Yet, these still suffer from a need for preincubation (Zoll et al. 2014), lack of selectivity, and a lack of activity against eukaryotic rhomboid enzymes (Tichá et al. 2017). For the reasons described above, these existing inhibitors are poor candidates to inhibit PfROM4, and thus a new targeting strategy is needed.

In this thesis, I demonstrate how I overcame many of these challenges. I began by deciphering the mechanistic basis of PfROM4’s atypical selectivity (Chapter II) and found a novel mechanism for achieving this selectivity. Next, I used that knowledge, along with other insights from our laboratory, to a design a selective, substrate-based inhibitor of this critical parasite enzyme (Chapter III). This compound, termed RiBn (rhomboid-inhibiting boronate), blocked invasion and cleared culture of blood-stage cultures of parasitemia.

Not only does this work establish for the first time that PfROM4’s activity is necessary and targetable for parasite invasion, it also provides a strategy for designing selective inhibitors for rhomboid enzymes from other disease-causing organisms.

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Figure 1.1: Malaria Lifecyle and Invasion Model

(A) The blood-stage lifecyle of P. falciparum is depicted. Malaria’s hallmark periodic fevers coincide with the synchronous egress of parasites from erythrocytes. Mature-stage parasites cause complications such as cerebral malaria by binding to endothelial cells and occluding the microvasculature. (B) Merozoite invasion into erythrocytes is depicted. Initial attachment is mediated by the MSP family of proteins. Following apical reorientation, the

EBL and Rh family of adhesins bind to their cognate receptors on the erythrocyte cell- membrane, thereby committing parasites to invasion. The AMA/RON complex forms the tight junction, an electron dense sieve that is connected to the parasite’s actin-myosin motor and mediates forced invasion of the erythrocyte. As invasion progresses, the MSP, AMA,

EBL, and Rh families of adhesins are shed from the parasite cell-surface, allowing the parasitophorous vacuole to successfully seal off at the posterior end of the parasite. (C)

PfSUB2, the first characterized ‘sheddase’, cleaves its substrates (MSP1 and AMA1) in their juxtamembrane segments. PfROM4, the putative intramembrane ‘sheddase’, cleaves its substrates (the EBL and Rh families) near the surface-exposed side of their transmembrane segments.

Figure 1.1 attachment MSP1 AB Malaria Blood-stage Lifecycle merozoite egress endothelial cell rhoptries merozoites erythrocyte

micronemes apical EBA/Rh schizont reorientation tight junction parasitophorous erythro- vaculoe cyte tropho- zoite invasion invasion

ring MSP1 EBA/Rh AMA/RON Blood vessel SUB2 ROM4?? capping and sealing time: <1 minute

C erythrocyte

chemical inhibition? selectivity? EBA-175 binding? MSP1 accessibility? kinetics?

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The malaria rhomboid PfROM4 exhibits atypical selectivity, cleaving critical parasite ligands but not canonical rhomboid substrates. In this chapter, I mapped the basis of

PfROM4’s atypical selectivity to the juxtamembrane sequence of substrates. Unexpectedly, I found that this recognition operates by steric exclusion – while PfROM4 can cleave a diversity of juxtamembrane sequences derived from parasites, it cannot cleave canonical substrates due to specific steric clashes between substrate and enzyme. Most notably, the enzyme was sensitive to only Ƣ-branched residues at the substrate P4 position: a single Val placed here was sufficient to abrogate cleavage. Next, I mapped this selectivity to a P4 interacting site on PfROM4 that when engineered onto a rhomboid homolog, endowed the ability to cleave PfROM4 substrates. Finally, I demonstrated that steric exclusion also drives the substrate selectivity of other rhomboid enzymes. Together, these results support a mechanism for achieving substrate selectivity in a membrane-immersed environment that is distinct from rules governing the selectivity of soluble proteases.

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After fifteen years of investigation, how rhomboid proteases recognize and process their substrates remains poorly understood, and represents an unresolved question. Unlike the case in soluble proteases, the field has not settled on a set of universal principles that predict if a particular transmembrane segment would be cleaved by rhomboid, hindering the search for novel substrates (Sinisa Urban 2010). Elucidating a mechanistic basis for substrate selectivity would not only provide important clues to the biological function of rhomboid proteases, but could also help to design specific and potent inhibitors (Drag and Salvesen

2010). Indeed, an understanding of the substrate selectivity of other proteases has been exploited to develop therapeutic agents for a variety of clinical indications including, but not limited to, hypertension, clotting disorders, HIV, HCV, and cancer (Turk 2006).

In biological systems, selectivity can be thought of in terms of the relative activity of a protease acting on alternative, competing substrates. It is quantitatively defined by a ratio of the maximum velocity of an enzyme to its Michaelis constant for a particular substrate, or kcat/KM (Hedstrom 2002). Because this selectivity constant does not depend on the concentration of either substrate or enzyme, it can be used to assess the substrate selectivity of various rhomboid enzymes in non-kinetic, physiologically relevant assays. These studies have painted a complicated, often contradictory picture of rhomboid protease substrate selectivity (Baxt et al. 2008; Baker, Wijetilaka, and Urban 2006; Riestra et al. 2015; Moin and

Urban 2012; Dickey et al. 2013; Strisovsky, Sharpe, and Freeman 2009; Sinisa Urban and

Freeman 2003; Akiyama and Maegawa 2007).

While the rules governing substrate selectivity differ between soluble and intramembrane proteases, the topic is much better understood in soluble proteases. Studies

(reviewed in Hedstrom 2002) in serine proteases have highlighted the importance of the

35 residues on the N-terminal side of the hydrolytic bond. These residues are termed P1-Pn with P1 being the residue on the N-terminal side of the scissile bond (those on the C- terminal side are similarly named P1’-Pn’) (Schechter and Berger 1967, Figure 2.1A). In chymotrypsin, trypsin, and elastase, the P1 residue determines selectivity: chymotrypsin prefers hydrophobic residues, trypsin cleaves after Arg/Lys, and elastase can only accommodate small aliphatic residues (Perona and Craik 1995). Moving outwards from the scissile bond, the addition of a amenable P3 residue increases catalysis by elastase by 100- fold (Stein et al. 1987). Finally, some proteases even act like specific nucleases: enterokinase requires the specific P5-P1 sequence D-D-D-D-K (Lu et al. 1999) for cleavage.

A widespread model of rhomboid substrate selectivity posits that, like serine proteases, rhomboid recognizes motifs surrounding the scissile bonds of substrates. This view is primarily based on a study in which the P5-P2’ residues of a Providencia stuartii rhomboid substrate, TatA, were comprehensively mutagenized and analyzed for in vitro cleavage by its physiological rhomboid enzyme, AarA (Strisovsky, Sharpe, and Freeman

2009). They found three significant position-dependent residue requirements: large, hydrophobic residues at the P4 position, small residues at P1 (confirming earlier data

(Akiyama and Maegawa 2007), discussed below), and a hydrophobic residue at P2’. A subset of these residue requirements was corroborated in other rhomboid substrate and enzyme pairings, both native and non-native. Finally, they used this ‘motif’ in a bioinformatics search to identify 13 additional putative substrates from P. stuartii, 5 of which were confirmed as substrates in the in vitro AraA cleavage assay. These P5-P2’ residue requirements have since formed the basis of ‘universal rhomboid recognition motif’ that has been employed – largely unsuccessfully - in the search for novel substrates of other rhomboid enzymes ((Riestra et al.

2015), unpublished data).

An alternative model, largely developed by our laboratory, disputes the notion that rhomboid relies on a specific ‘recognition motif’ to find and cleave its substrates. Its inception lies in the first investigation into substrate requirements for rhomboid cleavage where Sin Urban utilized a series of substrate and non-substrate chimeras to identify 7 residues at the top of Spitz’s transmembrane domain that were both necessary for Spitz cleavage and sufficient to induce cleavage of non-substrates such as TGF-ơ (Sinisa Urban and Freeman 2003). A more thorough analysis attributed cleavage to just two amino acids -

Gly-Ala - in the P5’-P6’ positions (precise cleavage site determined later (Moin and Urban

2012)). Though it was surprising that just two amino acids could mediate cleavage, the fact that they were helix destabilizing provided an intriguing explanation: they allowed the ơ-helix to unwind to access the active site for hydrolysis. This hypothesis was later confirmed via mutational analysis which demonstrated that the ơ-helix destabilizing residue proline could induce rhomboid cleavage of non-substrates when introduced into their transmembrane segments downstream of the scissile bond (Moin and Urban 2012). Thus, the presence of distal helix-breaking residues, not a specific sequence motif, was found to be the primary predictor of rhomboid cleavage.

The model that rhomboid cleavage depends on dynamics of the substrate has since been validated by our group and others. An independent investigation confirmed the necessity of downstream helix-breaking residues in the cleavage of additional rhomboid substrates (Akiyama and Maegawa 2007). The study also added one important requirement for the E. coli rhomboid enzyme GlpG: the preference for negatively charged residues in the

P1 and P1’ positions. While the later requirement is disputed for both GlpG and other rhomboid enzymes (Sinisa Urban 2010), the P1 and helix-breaking residue requirements have, to date, stood up to scrutiny.

Moin and Urban provided mechanistic insight into the role of helix-breaking residues in transmembrane segments by examining substrates and non-substrates for helicity via circular dichroism (Moin and Urban 2012). They found that the hydrophobic transmembrane segments of substrates tended to lose helicity in detergent whereas those of non-substrates remained helical. Reconstituting substrate transmembrane domains into membranes induced helicity, masking the instability. Additionally, they induced cleavage site shifts by introducing helix-breaking residues into distal positions within the transmembrane segment. These results reinforced two points: 1) As a consequence of helix-breaking residues, the transmembrane segments of rhomboid substrates are unstable, allowing them to unwind and access the active site. 2) Given that the cleavage site shifted easily, there are no strict residue requirements in the P5-P2’ sequence aside for a preference for small residues immediately flanking the scissile bond.

Finally, a kinetic analysis of various substrates and rhomboid enzymes in proteoliposomes revealed that differences in the selectivity constant, kcat/KM, between substrates and non-substrates was driven by kcat and not KM (Dickey et al. 2013). Even a substrate identified by Strisovsky, et al. as uncleavable (forming the basis for the recognition motif) actually had higher affinity (lower KM) for rhomboid than the WT substrate; slow relative cleavage was driven by a 100-fold decrease in kcat. Moreover, the measured KM was much higher than physiological concentrations of substrates within the membrane, indicating that affinity was unlikely to govern selectivity in rhomboid’s native environment.

Collectively, these data support the model in which the substrate selectivity of rhomboid enzymes is based the dynamics of substrate transmembrane domains and not affinity for substrates.

One prediction that emerges from this model is that rhomboid enzymes should be relatively non-selective: the dynamic properties that turn a transmembrane segment into a substrate for a rhomboid enzyme should make it a substrate for other rhomboid enzymes as well. In fact, this is what has been observed in various studies of rhomboid enzymes and their substrates (Sinisa Urban, Schlieper, and Freeman 2002; Kanaoka et al. 2005; Sinisa

Urban and Freeman 2003; Sinisa Urban and Wolfe 2005). The lone exceptions appear to be a handful of rhomboid enzymes from eukaryotic parasites (Baker, Wijetilaka, and Urban

2006; Baxt et al. 2008; Riestra et al. 2015). In a heterologous cleavage assay (Figure 2B), these enzymes are able to cleave parasite substrates such as EBA-175 and BAEBL (P. falciparum) but not canonical substrates like Spitz, whereas metazoan rhomboid display the opposite cleavage profile (Figure 2C)(Baxt et al. 2008; Baker, Wijetilaka, and Urban 2006; Riestra et al.

2015). On this basis, some parasitic rhomboid enzymes, including PfROM4 from P. falciparum, were deemed to exhibit atypical selectivity (Baker, Wijetilaka, and Urban 2006).

Despite ~10 years of investigation, the mechanistic basis of this atypical selectivity remains poorly understood and is difficult to reconcile with the model that transmembrane dynamics is the primary determinant for rhomboid cleavage. Defining the principles that govern this atypical selectivity therefore, would answer two major questions: 1) How is rhomboid selectivity achieved in a dynamics driven model? 2) Is the differential selectivity exhibited by parasitic rhomboid enzymes exploitable for the rational design of inhibitors?

Previous studies in PfROM4 have mapped its selectivity to the transmembrane segment of its substrates (Figure 2D): exchanging the transmembrane segment of BAEBL, a

PfROM4 substrate, for that of AMA1, a non-substrate, abrogated cleavage by PfROM4.

Conversely, substituting the transmembrane domain of AMA1 for that of BAEBL rendered the chimeric molecule cleavable by PfROM4 (Baker, Wijetilaka, and Urban 2006). Thus,

39 features within the transmembrane domain were thought to be sufficient to impart selectivity.

In this chapter, I will expand on these preliminary experiments to build a new model of rhomboid selectivity by answering three questions: 1) What are the requirements for

PfROM4 cleavage of native substrates? 2) What are the requirements for turning non-

PfROM4 substrates into substrates? 3) What is the enzymatic basis for these selectivity preferences?

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PfROM4 does not have strict P5-P2 residue requirements for cleavage

Given that PfROM4’s selectivity had been mapped to the transmembrane segment, I focused my initial experiments on P5-P2 residues of the PfROM4 substrate EBA-175, an important, essential merozoite adhesin (Camus and Hadley 1985; Duraisingh et al. 2003).

The P5-P2 region was selected for two reasons: 1) PfROM4 substrates uniquely featured aromatic residues (Phe, Tyr) in their P3-P2 positions (Figure 2.1D). 2) Residues in these positions (upstream of the scissile bond) are attractive targets for investigations into substrate selectivity both because they tend to be most important for cleavage by other proteases (Hedstrom 2002), and because they can help guide the development of substrate based inhibitors (Drag and Salvesen 2010). To test if such residues were necessary for cleavage by PfROM4, I exchanged the P5-P2 residues of EBA-175 (MPYY), a PfROM4 substrate, for those of two non-substrates - TgAMA1 (TALI, (Howell et al. 2005)) and Spitz

(MLEK, (S Urban, Lee, and Freeman 2001)) – and probed PfROM4’s ability to cleave these chimeras as well as the hydrophobic segment VLVV.

Contrary to my expectations, I found that PfROM4 was able to cleave both chimeras efficiently, indicating that EBA-175’s native P5-P2 residues were not a requirement for

40 cleavage (Figure 2.2A, Lanes 3-4). The Ƣ-branch heavy P5-P2 sequence VLVV was also cleaved, but not as efficiently. Even wholescale replacement of EBA-175’s transmembrane segment with that from Spitz did not abrogate cleavage, as would be expected if their respective transmembrane segments imparted reciprocal selectivity (Figure 2.2A, Lane 6).

Aside from VLVV, I mapped all cleavage sites via FLAG-immunoprecipitation followed by

MALDI-TOF analysis (Figure 2.1A).

Given that PfROM4 did not appear to require a specific P5-P2 motif or even transmembrane segment, I next designed a series of modifications to EBA-175’s TMD and juxtamembrane region with the goal of identifying a targeting motif that was necessary for

PfROM4 cleavage of EBA-175. These included increasing the length of transmembrane domain and incorporating various components from Spitz’s TMD and juxtamembrane segment. These were almost universally cleaved by PfROM4 (Table 1, highlighted in blue), indicating that this enzyme did not appear to have stringent transmembrane or juxtamembrane domain requirements for cleavage.

The only substrates that were not processed well by PfROM4 were those that included the sequence VLVV in the presumed P5-P2 positions (Table 1, highlighted in red).

No other enzyme appeared to be sensitive to this sequence (Table 1). Moreover, PfROM4 cleaved EBA-175 containing the Spitz-TM at a site downstream of the DmRho1 cleavage site (cleavage site depicted in Figure 2.2A), indicating that PfROM4 maintains a different sequence preference than other rhomboid enzymes even when cleaving the same substrate.

These results indicate that while PfROM4’s selectivity may be more complicated than originally anticipated, it still exhibits a differential selectivity that could be exploited if its underlying mechanism could be elucidated.

In summary, PfROM4 does not have strict P5-P2 residue requirements; significant amino acid diversity was tolerated in the P5-P2 residues (Figure 2.2A, Table 1). While these data contradict the idea of a universal recognition motif (Strisovsky, Sharpe, and Freeman

2009), it does not explain why PfROM4 cannot cleave other rhomboid substrates such as

Spitz and AMA1, especially given that it can cleave Spitz’s TMD. Thus, there must be a yet unidentified element that determines PfROM4’s ability to cleave Spitz.

In performing this analysis, I also noticed that seemingly simple changes were sufficient to turn EBA-175 into a substrate for metazoan enzymes (Table 1, data not shown).

For H. sapiens rhomboid 2, the substitution of EBA-175’s juxtamembrane residues for those of Spitz or extending the transmembrane length was sufficient to turn EBA-175 into a substrate that was cleaved at the native cleavage site. For DmRho1, in addition to Spitz’s juxtamembrane sequence, the P5-P2 residues from Spitz (MLEK) were also required for cleavage; either component alone was not sufficient to turn EBA-175 into a DmRho1 substrate. Thus, these metazoan enzymes also appear to have distinct mechanisms of substrate recognition.

Spitz is not conducive for cleavage by PfROM4

To determine if there was a targeting motif that was necessary and sufficient to turn

Spitz into a PfROM4 substrate, I engineered over 50 mutants of Spitz and probed them for cleavage by PfROM4 (Table 2). My initial hypothesis was that the P5-P2 residues from

EBA-175 would constitute a recognition motif (the aforementioned results notwithstanding), and induce cleavage by PfROM4 if placed into the appropriate position of Spitz. To test this model, I replaced the Spitz P5-P2 residues with EBA-175’s (MPYY) onto backgrounds with

TMD deletions of various lengths on account of EBA-175’s shorter transmembrane segment compared to Spitz (~16 vs 22 amino acids). While all constructs were cleaved by

DmRho1, Spitz’s physiological protease, none could be cleaved by PfROM4 (Figure 2.2B,

Table 2), indicating both that EBA-175’s P5-P2 residues were not sufficient to endow cleavage by PfROM4 and that Spitz’s P5-P2 residues were not necessary for DmRho1 cleavage. Combined, these results cast further doubt on the recognition motif model.

Even wholescale replacement of Spitz’s juxtamembrane and transmembrane segments with those from EBA-175 did not induce cleavage by PfROM4 (Figure 2.2C). All other rhomboid enzymes tested, parasitic or not, could cleave this chimera. Because many of these enzymes are active at the plasma membrane where PfROM4 is thought to be active, it is likely that this Spitz chimera is accessible to PfROM4 and a negative result represents a true lack of cleavage.

While our group had previously found that PfROM4’s atypical selectivity was mediated by the transmembrane domain (Baker, Wijetilaka, and Urban 2006), these results indicate that the ability to cleave Spitz is determined by features outside of the TMD: Spitz’s

TMD is cleavable when in EBA-175’s backbone (Figure 2.2A), but no mutation to Spitz’s

TMD - not even replacement with EBA-175’s TMD (Figure 2.2C) – rendered Spitz cleavable by PfROM4. Thus, it appeared that either Spitz’s C or N-terminal domain precluded cleavage by PfROM4. To distinguish between these possibilities, I generated a series of N and C-terminal chimeras between PfROM4 substrates and non-substrates.

Spitz’s ectodomain precludes PfROM4 cleavage

I began mapping selectivity determinants for PfROM4 cleavage of Spitz by replacing the N-terminal ectodomain of Spitz with that from EBA-175 (substrate) and AMA1 (non- substrate). Both ectodomains permitted PfROM4 cleavage of the Spitz TM and C-terminus

(Figure 2.2D, Table 3). While it is possible that EBA-175’s ectodomain provided positive affinity for PfROM4, thereby imparting selectivity, it is unlikely that AMA1’s ectodomain

43 could do so as well, because, unlike EBA-175, AMA1 is a not a PfROM4 substrate and does not share homology with EBA-175 (Figure 2.2E). Thus, the most likely explanation for these data is that Spitz’s ectodomain precluded cleavage by PfROM4.

A point that will be expanded upon in the next section is that cleavage was also dependent on the identity of the P5-P2 residues: those from AMA1 (KIII) were nonpermissive for cleavage, while those from Spitz (MLEK) allowed cleavage (Figure 2.2D).

To test if Spitz’s ectodomain indeed precluded PfROM4 cleavage, I replaced the N- terminal ectodomain of a PfROM4 cleavable mutant of AMA1 (see next section) with that from Spitz. AMA1 was chosen for the c-terminal because it was unlikely to impart specificity* for PfROM4. Replacing the ectodomain of the AMA1 mutant with that from

Spitz selectively reduced cleavage by PfROM4 but not by its ortholog from Toxoplasma gondii,

TgROM5 (Figure 2.2F, highlighted in red and green). Neither TgROM5 nor PfROM4 is likely to have positive affinity for the ectodomain of Spitz, a metazoan protein involved in growth signaling in multicellular organisms. Thus, the most plausible explanation for these data is that Spitz’s ectodomain is simply not-conducive for PfROM4 cleavage.

These experiments therefore highlight two requirements for PfROM4 cleavage of substrates: 1) A permissive ectodomain: Spitz’s ectodomain precludes PfROM4 cleavage, likely as a result of its proximity to the membrane (2.2G), which causes it to clash with the large Loop 1 of PfROM4. EBA-175 (native substrate) and AMA1 (non-substrate) contain longer juxtamembrane linkers to their non-homologous ectodomains and are thus permissive for PfROM4 cleavage. 2) The P5-P2 sequence: AMA1’s native P5-P2 sequence

(KIII) did not allow for cleavage whereas those from Spitz (MLEK) did (Figure 2.2D), indicating that PfROM4 has specific requirements for the identity of the P5-P2 residues that were not revealed by the previous analysis.

To elucidate this second requirement, I embarked on a series of experiments where I extensively mutagenized these residues in a variety of substrates and determined their effect on PfROM4 cleavage.

Ƣ-branched, hydrophobic residues in AMA1’s P4-P2 positions preclude PfROM4 cleavage

The ability for PfROM4 to cleave AMA1 appears to be mediated by residues of

AMA1’s TMD: AMA1 is cleavable when its TMD is replaced with that of a substrate (Baker,

Wijetilaka, and Urban 2006), and AMA1’s P5-P2 residues (KIII) are sufficient to abrogate cleavage of the Spitz chimera (Figure 2.2D). To understand the P5-P2 requirements for

PfROM4 cleavage of AMA1, I mutated these AMA1 residues to those from other PfROM4 substrates and non-substrates. Both MPYY (from EBA-175, substrate) and MLEK (from

Spitz, non-substrate) strongly induced cleavage by PfROM4 (Figure 2.3A). Additionally, mutating the P4-P2 III residues to AAA also permitted PfROM4 cleavage. Mass spectrometry analysis revealed that the MPYY chimera was cleaved at the native PfROM1 cleavage site, while the KAAA mutant was cleaved in multiple locations (Figure 2.3C), indicating that residues in these positions may not only the influence selectivity of PfROM4, but can also guide how it cleaves its substrates.

Whereas a diverse panel of amino acids at the top of the TMD induced cleavage of

AMA1 by PfROM4, hydrophobic and Ƣ-branched residue sequences such as TALI, VLVV and the native KIII were not cleaved efficiently (Figure 2.3A, red arrows). These results mirror those from EBA-175 where the Ƣ-branch residue heavy VLVV P5-P2 sequence was not cleaved as efficiently as other sequences (Figure 2.2A).

Individual P4-P2 amino acids strongly induce cleavage of AMA1 by PfROM4

To narrow this selectivity to a precise position, I mutated the P4-P2 residues to Ala,

Val and their counterparts from EBA-175 (P4-P2: PYY). A number of single amino acid mutations induced cleavage by PfROM4, including a P4 Pro, P3 Ala or Tyr, or a P2 Tyr

(Figure 2.3B). Mass spectrometry analysis confirmed that there were no cleavage site shifts for the residues examined (Figure 2.3C). While Ala, Pro and Tyr were all able to induce cleavage when introduced at various positions, the Ƣ-branched residue Val, like the native

Ile, was not permissive for PfROM4 cleavage.

Though single residue substitutions had a profound effect on the ability for PfROM4 to cleave of AMA1, this analysis did not identify a single, critical position that mediated

PfROM4 cleavage as substitutions at multiple positions induced cleavage. Given that AMA1 is a non-substrate, though, it is not likely to be recognized by PfROM4 as a substrate would be. Thus, I turned my focus to P5-P2 residues of EBA-175, the only genetically validated

PfROM4 substrate (O’Donnell et al. 2006).

PfROM4 is sensitive to the nature of EBA-175’s P4 residue

I began my exploration of EBA-175 by mutating its P4-P2 residues both in isolation and in combinations to Ile (from AMA1) and Ala. I found that the P4-P2 residues from

AMA1, III, abrogated PfROM4 cleavage entirely, an effect that could be explained by the presence of a single Ile at the P4 position (Figure 2.4A, red arrows); neither a P2 nor a P3 Ile had any effect on cleavage. Cleavage of other mutants did not differ significantly from WT

(Figure 2.4A, data not shown).

Given that PfROM4 was sensitive to mutation at the P4 position, I expanded the analysis at this position to include all 20 naturally-occurring residues (Figure 2.4B).

Surprisingly, most residues – positive, negative, large, aromatic, polar, hydrophobic – were cleaved efficiently. Some (Arg, Lys, Asp and Glu) were even preferred. Only those amino

46 acids that contain a CƢ branch (Ile, Val, Thr, red arrows) inhibited cleavage: EBA-175 containing a P4 Ile or Val was cleaved only ~20% of the WT Pro (quantification not shown).

These experiments demonstrate that 1) the P4 position of EBA-175 is a critical mediator of substrate selectivity and 2) PfROM4 does not recognize a particular motif.

Rather, it appeared that substrate selectivity is mediated by steric clashes: most chemical diversity is tolerated at the P4 position; it is the Ƣ-branched residues that preclude cleavage by PfROM4.

To investigate if such steric interactions governed the substrate selectivity of other rhomboid enzymes in addition to PfROM4, I performed a selectivity analysis on PfROM1, a rhomboid enzyme that is also expressed in blood-stage parasites.

The P4 residue acts as a selectivity switch for PfROM4 and PfROM1

PfROM1, the other active Plasmodium rhomboid enzyme , can cleave AMA1but not

EBA-175 (Baker, Wijetilaka, and Urban 2006). I interrogated its ability to cleave the panel of

EBA-175 mutants I had created for my PfROM4 substrate selectivity analysis. To my surprise, PfROM1 demonstrated selectivity that was almost perfectly reciprocal to PfROM4: it could efficiently cleave EBA-175 harboring P4 residues that inhibited PfROM4 cleavage

(Ile, Val and Thr) but not those that permitted cleavage by PfROM4 (Figure 2.4C, green arrows). I confirmed that these residues were in fact in the P4 position of EBA-175 when cleaved by PfROM1 by performing cleavage site analysis (Figure 2.4D). All constructs tested were cleaved at the native PfROM4 cleavage site.

An attractive model for achieving substrate selectivity emerges from these data: rhomboid proteases are inherently non-sequence specific. In a cell/organism where two rhomboid proteases are active, there needs to be a mechanism for imparting specify if

47 cleavage is regulated. In the case of PfROM4 and PfROM1, evolution achieved this by steric occlusion by Ƣ-branched residues.

Residues at end of rhomboid L1 loop of mediate steric clashes

I next sought to interrogate the enzymatic basis of these steric clashes at the P4 position. I began with the observation that a residue (F146) at the L1 loop from the E. coli rhomboid enzyme, GlpG, appears to interact with the P4 residue (Cho, Dickey, and Urban

2016; Zoll et al. 2014; Figure 2.5A), and may influence that enzyme’s preference for the P4 residue (Zoll et al. 2014). Guided by universally conserved residues (a structurally important

R, the oxyanion stabilizing H, and another H of unknown function), I constructed a structure-based alignment of various parasitic rhomboid enzymes, including PfROM1/4,

Trichomonas vaginalis rhomboid 1, and Toxoplasma gondii rhomboid 5 and found significant diversity at positions equivalent to EcGlpG 146 and 147 (Figure 2.5B).

I began by investigating if residues at these positions correlated with the differential substrate selectivity exhibited by the enzymes that harbored them (Figure 2.5C). While

TgROM5, TvROM1 and PfROM1 were all able to cleave a P4 Ƣ-branched isoleucine,

TvROM1 and PfROM1 were more active and have residues (Gly-Ser and Ala-Asn, respectively) that differ from PfROM4 (Gly-Gly) in this putative S4 binding pocket. To determine if this motif mediated selectivity, I created several enzyme L1 loop chimeras and probed their ability to cleave non-allowed residues. My expectation was that if selectivity was indeed governed by sterics, I could relieve the clash by introducing residues from enzymes able to tolerate those disallowed residues. The chimeric enzyme would gain the ability to cleave these residues without losing the ability to cleave tolerated residues. Alternatively, if selectivity was governed by positive interactions, the chimeric enzyme would gain the ability to cleave disallowed residues at the expense of previously allowed residues.

Simply installing a glycine at the first position endowed PfROM1 with the ability to cleave a variety of P4 residues (Figure 2.5E). The addition of a glycine at the second position further enhanced cleavage of these previously uncleavable substrates while retaining the ability to cleave the P4 isoleucine, indicating that the PfROM4 Gly-Gly motif relieved the steric clash rather than switching the selectivity of PfROM1. Importantly, this phenomenon was isolated to the P4/S4 interaction – the chimeric PfROM1 was unable to cleave a P2 variant that was cleaved by PfROM4 (Figure 2.5E, last column), indicating that the L1 loop mediates interactions specifically with the P4 residue.

Similarly, in the reciprocal experiment, introducing PfROM1’s L1 loop motif (Ala-

Gln) into PfROM4 allowed PfROM4 to cleave a P4 isoleucine (Figure 2.6B, 4th row) without affecting its ability to cleave proline, arginine or glutamate at the P4 position. Where

PfROM4 differed from PfROM1 was that the individual L1 loop amino acid substitutions

(alanine and glutamine, Figure 2.6B, 2nd, 3rd rows) had a minimal effect on the cleavage pattern. Still, like in PfROM1, these results supported the notion that substrate selectivity is guided by steric clashes between the enzyme and substrate; relieving this clash by introducing an accommodating L1 loop motif allows PfROM4 to cleave substrates harboring a P4 Ƣ-branched isoleucine.

Furthermore, substitution of a serine at the second position (G471S), not only imparted PfROM4 with the ability to cleave an isoleucine in the P4 position, it also appeared to enhance cleavage of all other substrates tested (Figure 2.6B, last row). When examined in parallel with the WT enzyme, G471S enhanced cleavage of WT-cleavable variants by 3-4 fold and the P4 isoleucine by 20-30 fold (Figure 2.6C, quantification shown 2.6D). A serine is naturally present at this position in both EcGlpG and TvROM1 (Figure 2.5B). In

EcGlpG, the serine appears to form a backbone hydrogen bond with TM2, potentially

49 stabilizing the confirmation of this Loop 1 motif (Cho, Dickey, and Urban 2016). Whether this residue plays a similar role in PfROM4 is unclear in the absence of structural data.

Nonetheless, this result provides further support for the notion that the L1 loop mediates steric interactions with the P4 residue of substrates.

In summary, these results support the hypothesis that steric clashes between enzyme and substrate mediate substrate selectivity in PfROM1 and PfROM4. As predicted by my model, the L1 loop chimeric enzymes gained the ability to cleave substrates containing disallowed residues without losing the ability to cleave those with allowed residues. That this effect was limited to the P4 position – the P2 selectivity remained intact in these experiments

– provided experimental evidence for the L1 loop/P4 interaction noted in crystal structures.

PfROM4 L1 loop variants can cleave TatA, a canonical rhomboid substrate

In initial investigations, PfROM4’s selectivity was characterized as atypical given that it could cleave substrates derived from parasites but not canonical rhomboid substrates

(Baker, Wijetilaka, and Urban 2006). In Figure 2.2, I demonstrated that one such canonical substrate (Spitz) was unable to be cleaved by PfROM4 due to a non-permissive ectodomain.

The Providencia stuartii rhomboid substrate TatA, however, was unable to be cleaved by

PfROM4 despite lacking an ectodomain entirely. It did, though, have an isoleucine in the P4 position. Thus, I asked if the PfROM4 L1 loop variants (G471S and AN) that could cleave a

P4 isoleucine in an EBA-175 backbone were also able to cleave TatA. As predicted, these chimeras gained the ability to cleave TatA (Figure 2.6E, green arrows) but not Spitz with its nonpermissive ectodomain (Figure 2.6F). These chimeras, however, were still unable to cleave AMA1’s TM segment (Figure 2.6G), indicating other steric mechanisms may be at work given the number of Ƣ-branched residues in the P4-P2 positions (III) of that transmembrane segment. That said, the fact that these residues also appear to govern the

S4/P4 interaction, and, as a consequence, cleavability, of another substrate demonstrates the robustness of the finding.

Steric clashes mediate substrate selectivity in other rhomboid enzymes

Separated by millions of years of evolution from other rhomboid enzymes, it is unclear if the described model of substrate selectivity is applicable outside of Plasmodium rhomboid enzymes. Thus, I investigated if substrate selectivity in other rhomboid enzymes was also governed by steric interactions. In an equivalent P4-P2 analysis as I performed with

PfROM4 (Figure 2.4A), the III motif abrogated cleavage by TvROM1, an effect that could be explained by the presence of a single isoleucine in the P2, rather than P4, position.

Similarly, in performing a separate analysis of Homo sapiens rhomboid-2 (hR2), I discovered that a number P3 mutations were strongly activating, turning EBA-175 into a hR2 substrate

(subset shown in Figure 2.7B). Therefore, it appears that the model of steric clash mediated by negatively clashing residues also applies to other rhomboid enzymes, albeit at differing sites.

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In this chapter, I have described a new model for how rhomboid enzymes achieve differential substrate selectivity: steric exclusion by negatively acting residues on both substrate and enzyme. My data overturn prevailing dogma in the field that rhomboid enzymes act like other site-specific proteases (Hedstrom 2002) and recognize their substrates by a specific motif upstream of the scissile bond (Strisovsky, Sharpe, and Freeman 2009;

Zoll et al. 2014). In generating this model, I created over 200 substrate and enzyme variants and found that rhomboid enzymes are tolerant of most residues in the P5-P2 residues of substrates, arguing against a positively acting recognition motif. Rather, they achieve

51 selectivity due to a limited number of negatively interacting residues; in the case of PfROM4, substrate P4 Ƣ-branched residues are not tolerated by L1 loop residues of the enzyme.

This model also explains how rhomboid enzymes can achieve selectivity despite having few non-backbone interactions between substrate-based peptide inhibitors and enzymes in crystal structures (Cho, Dickey, and Urban 2016; Zoll et al. 2014). In these structures, we can see how rhomboid is able to tolerate most residues – many R groups are oriented into solution – but because of the selection bias inherent in crystallization (ie we cannot co-crystallize peptides harboring non-tolerated residues), we can only hypothesize about the structural mechanism of negative selectivity. My L1 loop enzyme chimeras, however, provide powerful experimental evidence for the existence of such a mechanism.

This mechanism of substrate selectivity may provide one explanation for how multiple active rhomboid enzymes maintain differential substrate selectivity when active in the same cell/organism. For instance, Plasmodium spp., encode 8 different rhomboid proteins,

4 of which are essential (Lin et al. 2013), and 2 of which have been shown to be active

(Baker, Wijetilaka, and Urban 2006). The two active proteases, ROMs 1 and 4, are localized to different compartments (O’Donnell et al. 2006) and thus require a mechanism to prevent cleavage of each other’s substrates during trafficking. Steric exclusion endows these otherwise non-sequence specific enzymes with such a mechanism and would therefore be selected for.

While this analysis provides important answers to the question of how selectivity is achieved by rhomboid proteases, it does not easily lend itself to the design of substrate-based peptide inhibitors parenthetically. A new strategy, though, for designing such an inhibitor presented itself in my data: Some of the constructs that I generated appeared to be processed more efficiently than the WT substrate. Based on previous data from our lab (Cho, Dickey,

52 and Urban 2016), inhibitors designed from such sequences are more potent inhibitors of rhomboid enzymes. Thus, the genesis for the next chapter, designing a PfROM4 selective inhibitor, was based on the analysis in this chapter.

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DNA constructs

Open reading frames were cloned into pcDNA3.1 (Invitrogen) as N-terminal GFP- tagged/C-terminal Flag-tagged substrates or N-terminal 3xHA-tagged rhomboid proteases.

Expression was driven by a CMV promoter. EBA-175, AMA1, PfROM1/4 and TvROM1 were all recoded for human expression as described previously (Baker, Wijetilaka, and Urban

2006). Additionally, EBA-175 and AMA1 were cloned without their large ectodomains to both help with expression and streamline the analysis. Mutants were generated either via

QuikChange Site-Directed Mutagenesis (Agilent Genomics) or an inverse PCR strategy. All constructs were confirmed by DNA sequencing (Genewiz).

Heterologous HEK-cell based cleavage assay

Cleavage of these substrates and the activity of enzymes was determined in a slightly modified heterologous cleavage assay (Baker, Wijetilaka, and Urban 2006). Briefly,

HEK293T (ATCC CRL-11268) cells were seeded onto 12-well plates and ~80% confluent cells were co-transfected with the appropriate substrates and/or enzymes with 1.5 μL per well of X-tremeGENE-HP (Roche). Cells were transfected with a low amount of rhomboid enzyme: substrate:enzyme ~ 100-200:1 (1.25μg total DNA with 750ng bluescript DNA as filler). 18-hours post transfection, cells were washed once with 1mL of DMEM and conditioned in 400μL of serum-free DMEM (Sigma). For cells transfected with AMA1, the metalloprotease inhibitor BB-94 (20μM) was included to reduce spontaneous ectodomain shedding. Media containing the GFP-tagged N-terminal cleavage product and their

53 respective cells were harvested ~16-20-h later by re-suspension of lyophilized media and lysis of cells in Laemmli buffer. Protein from lysates and media were resolved on 4%-12% gradient Bis-Tris gels in MES running buffer (Life Technology), and transferred onto nitrocellulose membranes with a Trans-Blot (BioRad) semi-dry system. After blocking (Li-

COR Biosciences), membranes were probed with anti-GFP (Abcam, ab32146), anti-HA

(Roche, 3F10), or anti-FLAG (Sigma, F7424) primary antibodies followed by anti-rabbit/rat secondary antibodies conjugated to infrared fluorophores (Li-COR Biosciences) and imaged on an Odyssey infrared scanner (Li-COR Biosciences). Two-color quantitative blots were converted to greyscale.

Cleavage site analysis

To determine the cleavage site of rhomboid substrates, HEK293T cells were transfected in 6-well plates as described above, but with a higher amount of enzyme: substrate:enzyme ~50:1 and DNA (2.5μg total). 18-hours post-transfection, cells were washed 1x with DMEM conditioned with 1mL of serum-free media (DMEM). ~24 hours later, cells were resuspended in DMEM, pooled, pelleted (1000g x 10 minutes), and lysed in

500μL RIPA buffer in the presence of a protease inhibitor cocktail (Complete Tablets,

Roche). After homogenization in a cup horn sonicator (Branson) and filtration through a loading tip, samples were subjected to anti-FLAG immunopurification (Sigma). Briefly,

3xwashed anti-FLAG beads were incubated with samples for 1-1.5hrs at 4C. Agarose beads were washed 2x with RIPA buffer and detergent was removed with 2x ddH20. Bound peptide was eluted with a 4:4:1 mix of H2O:Isopropanol:Formic acid. Eluent was spotted onto a sinapinic acid matrix, and analyzed by MALDI-TOF mass spectrometry on a standards calibrated Voyager DE Instrument (AB SCIEX) as previously described (Moin

54 and Urban 2012). Resultant spectra were analyzed and plotted in the R environment with aid of the Maldiquant package (Gibb and Strimmer 2012).

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I thank Sin Urban for both designing the project and teaching me how to perform basic molecular biology techniques, including the heterologous cleavage assay. I thank Syed

Moin for teaching me how to perform the cleavage site analysis. I thank all members of the lab, particularly Seth Dickey, for insightful discussion and advice. I performed and analyzed all experiments described in this chapter.

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Figure 2.1: PfROM4 exhibits atypical specificity

(A) Substrate/Protease nomenclature is depicted. Beginning at the scissile bond, substrate residues are numbered P1-PX in the direction of the N-terminus and P1’-PX’ in the direction of the C-terminus. Their corresponding enzyme residues are termed SX’-SX. In many proteases, substrate specificity is governed by the P4-P1 residues. (B) For cleavage analysis, N-terminal GFP and C-terminal FLAG-tagged substrates and N-terminal 3xHA tagged rhomboid enzyme were co-transfected into HEK cells. GFP or FLAG-tagged cleavage products were detected by western blot analysis of the media and lysates, respectively. FL refers to the full-length substrate as expressed in cells. The N-terminal GFP cleavage product is released into the media upon rhomboid processing. The C-terminal

FLAG cleavage product is retained in the cells. To determine the cleavage site, FLAG-tagged

C-termini were subjected to immunopurifcation followed by MALDI-TOF analysis. (C)

PfROM4 exhibits atypical specificity, cleaving the Plasmodium adhesins EBA-175 and

BAEBL but not the canonical rhomboid substrate, Spitz. While the other parasitic rhomboid enzymes (TvROM1 and TgROM5) could also cleave EBA-175 and BAEBL, metazoan enzymes cleaved only Spitz. (D) The transmembrane segments of non-PfROM4 substrates

(AMA1 and Spitz) are depicted above those of substrates (EBA-175 and BAEBL). The latter are notable for the aromatic residues in P2 and P3 while AMA1 features three isoleucines in positions P4-P2.

Figure 2.2: Spitz’s ectodomain precludes PfROM4 cleavage

(A) The P5-P2 residues of EBA-175 were replaced with MLEK (from Spitz), TALI

(TgAMA1), and VLVV. Additionally, EBA-175’s whole transmembrane segment was replaced with that from Spitz (last column). Of these, only VLVV was cleaved inefficiently.

The cleavage sites were mapped and aligned as indicated on right. (B) Various modifications to Spitz, including replacing its P5-P2 motif with that from EBA-175 (MPYY), and truncating its transmembrane domain did not convert it into a PfROM4 substrate. (C) A construct, depicted on left, featuring wholescale replacement of Spitz’s transmembrane domain with that from EBA-175 was cleaved by all rhomboid enzymes tested except for

PfROM4 (2nd column). (D) As depicted (left) a chimera of AMA1’s ectodomain attached to

Spitz’s transmembrane domain with the junction at P5 (retaining Spitz’s native MLEK) was cleaved efficiently by PfROM4. A chimera with the junction was placed at P1 (P5-P2: KIII from AMA1, left), however, was not a PfROM4 substrate. (E) A chimera containing

AMA1’s ectodomain with the distal ~40 residues of EBA-175 was cleaved efficiently. The junction was at P5, retaining EBA-175’s native P5-P2 residues. (F) AMA1 containing EBA-

175’s P5-P2 residues is cleaved efficiently by all rhomboid enzymes aside from DmRho1

(top panel of western). When Spitz’s ectodomain is transplanted onto this construct (bottom panel), it abrogates cleavage by PfROM4 (red box) but not TgROM5 (green box), a close

60 homolog. (G) A model depicting the length of the extracellular linker before the ectodomain for AMA1, Spitz, and EBA-175. Spitz’s linker is relatively short and may preclude cleavage by the relatively bulky PfROM4.

Figure 2.3: The AMA1 P5-P2 residues govern PfROM4 cleavage

(A) A variety of P5-P2 sequences (MPYY, MLEK, KAAA) permitted PfROM4 cleavage whereas those featuring hydrophobic and Ƣ-branched residues (KIII, TALI, VLVV) were not PfROM4 substrates. (B) Individual P4-P2 amino acid analysis revealed that substituting many residues in the P4 and P3 positions of AMA1 permitted PfROM4 cleavage. The IIY construct (starred) had non-PfROM4 mediated release into the media (data not shown), presumably by an endogenous rhomboid enzyme. (C) Cleavage site analysis of PfROM4 cleavable constructs revealed that PfROM4 was able to tolerate a diversity of residues in the

P4-P2 residues of AMA1 substrates.

Figure 2.4: Exclusion by P4 residues mediates the differential specificity of PfROM1 and PfROM4

(A) Transplanting the P4-P2 motif from AMA1 (III) into EBA-175 abrogates PfROM4 cleavage (right arrow), an effect that was mediated by a single isoleucine in the P4 position

(left arrow). (B) Cleavage analysis of all 20 naturally occurring amino acids at the P4 position revealed that most were cleaved efficiently, some even more efficiently than the WT proline.

The Ƣ-branched isoleucine, valine, and threonine (red arrows), however, hindered PfROM4 cleavage. This analysis was performed with high and low (left and right panel, respectively) amounts of PfROM4 to account for the differential cleavage of hydrophobic (left) and non- hydrophobic residues (right). (C) PfROM4 (top) and PfROM1 (bottom) cleavage of a representative panel of P4 residue variants is shown. The Ƣ-branched residues isoleucine, valine, and threonine were efficient substrates for PfROM1 but not PfROM4. (D) Cleavage

61 site analysis of the P4 variants revealed that PfROM1 and PfROM4 both cleaved EBA-175 variants at the same, native, cleavage site.

Figure 2.5: The L1 Loop forms the S4 binding pocket and mediates steric interactions with P4 residues

(A) Crystal structure depicting the L1 loop of EcGlpG interacting with the P4 residue of a bound substrate based inhibitor (Cho, Dickey, and Urban 2016). The distal residues (F146 and S147) form the S4 binding pocket (right). (B) Structure based alignment of the L1 loop region from various rhomboid enzymes. The alignment is anchored by the universally conserved structural arginine as well as the oxyanion-stabilizing histidine. Residues corresponding to positions 146 and 147 on EcGlpG are highlighted in green. (C) A panel of parasitic rhomboid enzymes were analyzed for their ability to cleave EBA-175 P4 residue variants. TvROM1 and PfROM1 cleaved the P4 isoleucine construct efficiently (green arrows). Neither cleaved EBA-175 when an isoleucine was placed in the P2 position (last column). (D) Structure based alignment of the L1 loop region highlighting PfROM1 Æ

PfROM4 chimeras at positions corresponding to GlpG residues 146 and 147. (E) PfROM1

Æ PfROM4 L1 loop chimeras could cleave previously non-tolerated P4 residues (green arrows). They were unable to cleave the P2 isoleucine, indicating that this effect was specific to the P4 residue.

Figure 2.6: PfROM4 L1 loop chimeras confer the ability to cleave Ƣ-branched P4 residues

(A) Structure based alignment of the L1 loop region highlighting PfROM4 Æ PfROM1,

TvROM1 chimeras at positions corresponding to GlpG residues 146 and 147. (B) PfROM4

L1 loop chimeras (AN, G471S, green arrows) gained the ability to cleave a P4 Ƣ-branched isoleucine without losing the ability to cleave the P2 isoleucine. Additionally, G471S

62 appeared to enhance cleavage of all substrates. (C) Parallel analysis of the WT and G471S

PfROM4 enzymes. At equivalent levels of enzyme and substrate expression, the G471S variant exhibited greater activity against all substrates tested. (D) Quantification of the western blot in (C). Cleavage was normalized for enzyme and substrate levels (depicted in orange). Except for the P4 isoleucine (20-30 fold more active), G471S was ~3 fold more active than the WT enzyme. (E) Cleavage of a bacterial rhomboid substrate, TatA, by the

G471S and AN variants of PfROM4 (green arrows). Neither variant cleaved Spitz (F) or

BAEBL-AMAtm (G)

Figure 2.7: The steric exclusion mechanism governs rhomboid specificity

(A) TvROM1 cleavage assay is displayed as a two-color western. TvROM1 is unable to tolerate an isoleucine in the P2 position (red arrows). (B) P3 variants of EBA-175 convert it into a hR2 substrate (green arrows). (C) A model summarizing how the steric exclusion mechanism works to determine rhomboid enzyme specificity. Top left: Spitz’s ectodomain does not allow for cleavage of that substrate even though its TMD is permissive for cleavage

(green star). Top right: PfROM4 is unable to cleave Ƣ-branched residues in the P4 positions of substrates (indicated in red letters) but is otherwise able tolerate a breadth of residue chemistry at that position. Also depicted is the S4 binding site at the end of the L1 loop on the enzyme that mediates the steric clash. Middle row: PfROM1 can tolerate Ƣ-branched but not other residues at the P4 positions of substrates. Bottom: Chemical structure of a substrate highlighting how a P4 Ƣ-branched residue may clash with residues of the S4 pocket.

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FOHDYDJH Į+$ site +

LQVLGH 4874.7 FDWDO\WLFUHVLGXHV ,3 ◀ )/$*& 80 100 3 3 3 3 3 3¶3¶ 1+$ 0$/',72) 03<<$*$ 7UDQVLHQW+(.&HOO7UDQVIHFWLRQ 4875.7 Relative Intensity 0204060 4000 5000 6000 7000 8000 Mass (m/z) C Enzyme: Parasitic

- 3I5 TvR1 hR2 TgR5 'P5 Metazoan (%$ (%$ 3I5P. falicaprum ROM4 CleaveG TvR1 T. vaginalis ROM1 %$(%/ %$(%/ hR2 H. sapiens ROM2 DGKHVLQV &OHDYHG Plasmodium TgR5 T. gondii ROM5 Spitz 'P5 D. melanogaster5KR Spitz CleaveG

(*) Į*)3 MeGia Signaling D Selectivity? Small residues Cleavage Site Enzyme: Substrate: ź Helix breaking residues P4P3 P2 P1 P1’ DmRho1: Spitz L E K A S I A S G A M C A LVF MLF V C L A F Y L PfROM1: AMA1 IIIA SSAAV A VLA T ILMVY L Y EBA-175 P YYA G A G VLF IILVILG A S ~~~ PfROM4: BAEBL P Y F AAGG ILVVIVLLLSSA S ~~

Transmembrane Segment

64 Figure 2.2

A EBA-175

P5-P2: Cleavage Site VLVV SPItm MPYY MPYY MLEK TALI PfR4: -++ ++ + P5 P4P3 P2 P1 P1’ Mapped? EBA EBA MPYY (WT) M P YYA G 9 Cleaved Į*)3 media EBA MLEK MLE K A G 9 EBA TALI T A LIA G 9 EBA FL EBA VLVV VLVVA G Į*)3 EBA SPItm K A S I A S 9 DmR1 lysates cleavage site PfR4 HA Į

B Spitz P5-P2: MPYY C Spitz EBA-TM FL ǻ ǻ Enzyme: - PfR4 TvR1 hR2 DmR1 Enzyme: TgR5 Spitz - PfR4 DmR1 - PfR4 DmR1 - PfR4 DmR1 Cleaved media Į*)3 Spitz Spi Spitz FL

Cleaved media Į*)3 Spitz Į*)3

EBA Cleaved Spitz FL Spi Į*)3

N-Spi-EBAtm-Spi-C lysates PfR4 HA 49 kDa Į lysates HA

Į 37 kDa DmR1

D E AMA1 Ectodomain/ N-AMA1 Ectodomain/SpiTM-C (%$70&)/$* P5-P2 : KIII MLEK Residues PfR4:

PfR4: - wt mut - wt mut - wt mut

AMA Cleaved KIII/ Į*)3 media Į*)3 AMA Ecto media

MLEK Cleaved AMA AMA Ecto EBA AMA/ Spi Į*)3 AMA/ Į*)3 SpiTM EBA

N-AMA-SpiTM-C lysates lysates Į+$

N-AMA-EBATM-C PfR4 PfR4 Į+$

AMA1 P5-P2: MPYY F G AMA1 EBA-175

Enzyme: Spitz - PfR4 TvR1 hR2 TgR5 DmR1 AMA Spi AMA Ecto Ectodomain - MPYY TM Distance ~100aa ~600aa

AMA AMA ~20aa PfR4 SPI Ecto N-AMA-MPYY-AMA-C Į*)3PHGLD N-Spi-MPYY-AMA-C

65 Figure 2.3

A AMA1 YY 33 P5 P4P3 P2 P1 P1’ KIII (WT) 03 KAAA MLEK TALI VLVV 3I5 +++ +++ AMA KIII (WT) K IIII I I A S AMA AMA MPYY (EBA) M P YYA S * Cleaved media Į*)3 AMA MLEK (Spi) MLE K A S AMA TALI (TgAMA) T A LLII A S $0$)/ AMA KAAA K AAAAAA A S

Į*)3 * AMA VLVV VLVVA S lysates 3I5 HA

Į * cleavage site mapped

P4P3 P2 P1 P1’ B AMA1 AMA1 III (WT) IIIA S AMA1 AAA AAAAS* II 33 III AAA AII VII 3 IAI IVI IYI IIA IIV IIY (WT) AMA1 AII A IIA S 3I5 +++++++++++ AMA1 VII VI IA S AMA Cleaved * AMA1 PII P IIA S Į*)3 media AMA1 IAI I A I A S $0$)/ AMA1 IVI IVIA S Į*)3 AMA1 IYI I Y I A S

lysates * 3I5 HA AMA1 IIA IIAAS Į AMA1 IIV IIVA S QRQ3I520PHGLDWHGFOHDYDJH AMA1 IIY IIY A S* C * cleavage site mapped

AMA KAAA AMA M3YY AMA IYI 9410 967 971

9813 967 Relative Intensity Relative Intensity Relative Intensity Relative 0 40 60 80 100 0406080100 0406080100 0 0 0 9000 900 10000 1000 9000 900 10000 1000 9000 900 10000 1000 Mass (m/z) Mass (m/z) Mass (m/z) 3 33 3 3 3¶ 3¶ 3 33 3 3 3¶ 3¶ N M K AAAASS M P YYA SS K I Y I A S S 9813 9411

66 Figure 2.4

A EBA-175 P4P3 P2 P1 P1’ EBA PYY (WT) P YYA G P4-P2: AYY IYY PAY PIY PYA PYI PAA III PYY (WT) EBA AYY A ... . PfR4: ++++++++ + EBA IYY I ... . EBA EBA PAY . A . . . media Cleaved Į*)3 EBA PIY . I . . . EBA PYA . . A. . EBA PYI . . I . . (%$)/

Į*)3 lysates EBA PAA . AA . . EBA III III. .

*)3(BA175 B 2 ng PfR4 .5 ng PfR4 P4: PYW* CMA IV/) P TSNQ RK H DE

EBA Į*)3 Cleaved media

(%$)/ Į*)3 lysates

Cȕ-branched amino acids

Ile Val Thr C EBA-175 EBA-175 AMA1 P4: PDAIV/)TSRK E PfR4 P4 PfR1 P4 P I Y I Į*)3 Media Y I A A

PfR1 PfR4 P1 P1

D EBA MIYY PfR1 EBA MTYY PfR4 EBA MTYY PfR1 4878 4880 4879 P4 P/I/T Y Y

A PfR4 Relative Intensity Relative Intensity Relative Relative Intensity Relative G PfR1 P1’ 020406080100 020406080100 020406080100 4000 4500 5000 5500 6000 4000 4500 5000 5500 6000 4000 4500 5000 5500 6000 Mass (m/z) Mass (m/z) Mass (m/z)

67 Figure 2.5 P4/S4 interaction A Peptide-CHO P4 Val

L1 loop F146 End

S147

B C

EBA

Loop 1GlpG146 TM2 GlpG147 P4-P2: PYY IYY RYY EYY PYA PYI ...... GlpG R Y F T H A LMH F S LMH PfR4 WT PfROM4 R L F WS M Y L H GGF MH PfROM1 R LILP I F L H A N I F H TgR5 TvROM1 R L F T Y M F L H G S W I H TvR1 TgR5 R VVWG M F L H GGWMH rhomboid conserved anchoring residues PfR1 WT P4 interacting residues ĮGFP Media

D E EBA

Loop 1GlpG146 TM2 GlpG147 P4-P2: ...... PYY IYY RYY EYY PYA PYI GlpG R Y F T H A LMH F S LMH PfR4 WT PfROM4 R L F WS M Y L H GGF MH PfROM1 WT R LILP I F L H A N I F H PfR1 WT PfROM1 A144G R LILP I F L H G N I F H PfROM1 N145G R LILP I F L H A G I F H PfR1 A144G PfROM1 GG R LILP I F L H GG I F H

conserved anchoring residues rhomboid PfR1 N145G P4 interacting residues PfR1 GG ĮGFP Media

68 Figure 2.6

Loop 1GlpG146 TM2 GlpG147 ...... EBA

P4-P2: PYY IYY RYY EYY PYA PYI GlpG R Y F T H A LMH F S LMH PfROM4 R L F WS M Y L H GGF MH WT R LILP I F L H A N I F H PfROM1 WT G470A PfROM4 G470A R LILP I F L H A G I F H PfROM4 G471N R LILP I F L H G N I F H G471N R LILP I F L H A N I F H PfR4 PfROM4 AN AN PfROM4 G471S R LILP I F L H G S I F H G471S conserved anchoring residues ĮGFP Media P4 interacting residues

G471S/WT Cleavage CDPfR4 WT PfR4 G471S 35 EBA 30

P4-P2: PYY IYY RYY EYY PYA PYI PYY IYY RYY EYY PYA PYI 25 EBA Cleaved GFP 20 Į Media 15

EBA FL GFP

Į 10 Lysates PfR4 PfR4/EBA-175) WT 5 HA Cleavage (normalized to Į 0 G471S activity is ~3-4 fold higher than PYY (WT) IYY RYY EYY PYA PYI Raw Normalized PfROM4 WT

GFP-Baebl-AMAtm E GFP-TatA-FLAG FG GFP-Spitz-FLAG - - DmR1 PfR4 G471S PfR4 WT AN PfR4 PfR1 PfR4 G471S PfR4 WT AN PfR4 - DmR1 PfR4 G471S PfR4 WT AN PfR4 Baebl/AMA Cleaved

TatA Į*)3 Cleaved Spitz Media Į*)3 Media Cleaved Į*)3 Media

Baebl/AMA Į*)3 TatA Cleaved Cleaved PfR4 Į)/$* Spitz PfR4 Cleaved Į*)3 Lysates Lysates DmR1 Į+$ Spitz Į+$

Lysates Cleaved Į)/$* PfR1

69 Figure 2.7

EBA-175 + TvROM1 EBA: sP3R wt P4-P2: P3R sEBA PYY PIY PAY PYI PYA PII PAA III IYY AYY PfR4 EBA Į*)3 Media Cleaved rhomboid hR2

Į*)P media Į*)3 (%$)/ Lysates Į+$ TvR1

C EBA-175 AMA1 Spitz PfR4 P4 P4 S4 L1 P A L S ... I V T I A Y E L S I A + I A Y K I A I A P2 P2

AMA1 EBA-175

PfR1 P4 S P4 4 I L1 P I V T I Y I A Y + Y P2 P2 PfROM4

sterics L1 loop R S scissile P 4 S2 bond S1' 4 P P ' O O 2 O 1 O ȕ + + + N N N N- N N N 1+ -C + + + O O O P3 P1 P2' S3 S1 S2'

70 Table 1. Most EBA-175 variants can be cleaved by PfROM4

Name N-term JxtMem P5-P2 TM C-term DmR1 PfR4 hR2 TgR5 TvR1 Figure EBA WT WT WT WT WT WT -+-++2.2A P5-P2 MLEK (Spitz) WT WT MLEK WT WT +/- + + 2.2A P5-P2 TALI (TgAMA) WT WT TALI WT WT +/- + + 2.2A P5-P2 VLVV WT WT VLVV WT WT -+/-++ 2.2A P3-P2 VV WT WT MPVV WT WT -+++ P5-P4 VL WT WT VLEK WT WT -+-++ WT TM+31 WT WT WT WT+3 WT -+-++ WT TM+41 WT WT WT WT+4 WT -++++ WT TM+51 WT WT WT WT+5 WT -+++- WT TM+61 WT WT WT WT+6 WT -+++- Spitz-TM11 WT WT MLEK Spitz11 WT +/- + + + Spitz-TM11 IA2 WT WT MLEK Spitz11 I->A WT +/- + + + Spitz-TM17 WT WT MLEK Spitz17 WT +/-++++/- Spitz-TM WT WT MLEK Spitz WT +++++2.2A Spitz-TM ǻ2 WT WT MLEK Spitz ǻ2WT +++++ Spitz JxtMem WT Spitz WT WT WT -++++ Spitz JxtMem+MLEK WT Spitz MLEK WT WT +++++ Spitz JxtMem+TM17 ǻ2 WT Spitz MLEK Spitz17 ǻ2WT +++++ Spitz JxtMem+TM WT Spitz MLEK Spitz WT +++++ Spitz JxtMem+TM VLVV WT Spitz VLVV Spitz WT ++/-++ + Spitz JxtMem+TM17 ǻ2 VLVV WT Spitz VLVV Spitz17 ǻ2WT ++/-++ + Spitz JxtMem+TM17 ǻ2 VV WT Spitz MLVV Spitz17 ǻ2WT +++++ Spitz JxtMem+TM17 ǻ2 VL WT Spitz VLEK Spitz17 ǻ2WT +++++ Spitz JxtMem+TM IA2 WT Spitz MLEK Spitz I->A WT +++ + Spitz JxtMem+TM VLVV IA3 WT Spitz VLVV Spitz I->A WT +++++ Spitz JxtMem+TM17 IA2 WT Spitz MLEK Spitz17 I->A WT +++++ Spitz JxtMem+TM17 VLVV IA3 WT Spitz VLVV Spitz17 I->A WT +++++ 1 hydrophobic residues were added to the middle of the transmembrane segment to match the length of Spitz 2 this is the equivilant mutation to I144A in Spitz. In this context, the mutation allows for PfROM4 cleavage site shifts 3 VLVV is not cleaved well in the P5-P2 residues of the EBA/SpitzTM chimera. The I144A equivilant mutation rescues this cleavage defect

71 Table 2. Spitz cannot be modified into a PfROM4 substrate

Name N-term JxtMem P5-P2 TM C-term DmR1 PfR4 hR2 TgR5 TvR1 Figure Spitz WT WT WT WT WT WT +-++-2.1C P5-P2 IPYF (Baebl) WT WT IPYF WT WT +- - P5-P2 MPYY (EBA) WT WT MPYY WT WT +- - 2.2B WT I144A WT WT WT I144A WT +- + IPYF I144A WT WT IPYF I144A WT +- + MPYY I144A WT WT MPYY I144A WT +- + P1-P2' PYF WT WT WT P1-P2' PYF WT +-++- A142P WT WT WT A142P WT +/- - + - - WT TMǻ4 WT WT WT ǻ4WT+- WT TMǻ5 WT WT WT ǻ5WT+- WT TMǻ6 WT WT WT ǻ6WT+- + WT TMǻ7 WT WT WT ǻ7WT+- + WT TMǻ8 WT WT WT ǻ8WT+- - WT TMǻ9 WT WT WT ǻ9WT+- - WT TMǻ10 WT WT WT ǻ10 WT +- IPYF TMǻ4 WT WT IPYF ǻ4WT+- IPYF TMǻ5 WT WT IPYF ǻ5WT+- IPYF TMǻ6 WT WT IPYF ǻ6WT-- - IPYF TMǻ7 WT WT IPYF ǻ7WT-- - IPYF TMǻ8 WT WT IPYF ǻ8WT-- - IPYF TMǻ9 WT WT IPYF ǻ9WT-- - IPYF TMǻ10 WT WT IPYF ǻ10 WT -- MPYY TMǻ4 WT WT MPYY ǻ4WT+- 2.2B MPYY TMǻ5 WT WT MPYY ǻ5WT+- 2.2B MPYY TMǻ6 WT WT MPYY ǻ6WT+- + MPYY TMǻ7 WT WT MPYY ǻ7WT+- + MPYY TMǻ8 WT WT MPYY ǻ8WT-- - MPYY TMǻ9 WT WT MPYY ǻ9WT-- - MPYY TMǻ10 WT WT MPYY ǻ10 WT -- BAEBL-TM WT WT WT BAEBL WT --+++ EBA-TM WT WT MPYY EBA WT +-+-+/- EBA JxtMem+TM WT EBA MPYY EBA WT +-+++2.2C EBA JxtMem WT EBA WT WT WT +-++- EBA JxtMem TMǻ6 WT EBA WT ǻ6WT+-+-- MPYY + EBA JxtMem WT EBA MPYY WT WT +-++- MPYY ǻ6 +EBA JxtMem WT EBA MPYY ǻ6WT+-+-- IPYF + BAEBL JxtMem WT Baebl IPYF WT WT +-++- JxtMem duplication1 WT 2xSpitz WT WT WT +-++- JxtMem duplication TMǻ61 WT 2xSpitz WT ǻ6WT+-+++ C141S2 C141S WT WT WT WT +- C141S JxtMem dup2 C141S 2xSpitz WT WT WT +- C141S EBA JxtMem2 C141S EBA WT WT WT +- C141S EBA JxtMem+TM2 C141S EBA MPYY EBA WT +- ǻEGF3 ǻEGF WT WT WT WT +- +/- ǻGFP ǻEGF4 ǻGFP,EGF WT WT WT WT +- ǻGFP ǻEGF MPYY4 ǻGFP,EGF WT MPYY WT WT +/- - ǻGFP ǻEGF IPYF4 ǻGFP,EGF WT IPYF WT WT +/- - ǻGFP ǻEGF EBA-TM4 ǻGFP,EGF WT MPYY EBA WT +/- - ǻGFP ǻEGF BAEBL-TM4 ǻGFP,EGF WT IPYF BAEBL WT +/- - 1 increases the spacer between the membrane and the EGF-domain from ~15 to ~30 aa 2 disrupts the terminal disulfide bond in the EGF domain, allowing it to partially unfold 3 attaches the GFP domain to the juxtamembrane linker of Spitz 4 removes both GFP and EGF domains entirely. Cleavage analyzed by generation of C-terminal flag tagged cleavage product

72 Table 3. Chimeric substrates highlight importance of P5-P2 motif

Name Junction N-term P5-P2 TM C-term DmR1 PfR4 hR2 TgR5 TvR1 Figure EBA/Spitz P1 EBA MPYY Spitz Spitz +/- + + +/- - EBA/Spitz P5 EBA MLEK Spitz Spitz - AMA/Spitz P1 AMA KIII Spitz Spitz - 2.2D AMA/Spitz P5 AMA MLEK Spitz Spitz + 2.2D Spitz/EBA1 P5 Spitz MPYY EBA EBA ----- Spi/AMA2 P5 Spitz MPYY AMA AMA --+++2.2F Spi/AMA P5 Spitz KIII AMA AMA --++- EBA/AMA2 P1 EBA MPYY AMA AMA +/- + +/- + + AMA/EBA P5 AMA MPYY EBA EBA + 2.2E EBA/AMA P5 EBA KIII AMA AMA +/- 1 this construct is not cleaved by any enzyme, indicating a possible trafficing error 2 combined, these constructs indicate that Spitz’s ectodomain selectivly inhibits PfROM4 cleavage as no other enzyme is affected by the presence of the ectodomain

Table 4. AMA1’s P5-P2 motif precludes PfROM4 cleavage

Name N-term P5-P2 TM C-term DmR1 PfR4 hR2 TgR5 TvR1 Figure AMA WT WT KIII (WT) WT WT --+/-+-2.3A P5-P2 MLEK (Spitz) WT MLEK WT WT -++++2.3A P5-P2 TALI (TgAMA) WT TALI WT WT --+/-+-2.3A P5-P2 VLVV WT VLVV WT WT --++-2.3A P5-P2 KAAA WT KAAA WT WT -++++2.3A P5-P2 MPYY (EBA) WT MPYY WT WT -++++2.3A

Table 5. AMA1 P5-P2 point mutations induce PfROM4 cleavage

Name N-term JxtMem P5-P2 TM C-term PfR4 PfR1 Figure AMA WT WT WT WT WT WT -+/- 2.3B KAYY WT WT KAYY WT WT ++ KVYY WT WT KVYY WT WT ++ KIYY WT WT KIYY WT WT ++ KAII WT WT KAII WT WT -- 2.3B KVII WT WT KVII WT WT -- 2.3B KPII WT WT KPII WT WT +/- + 2.3B KIAI WT WT KIAI WT WT +/- +/- 2.3B KIVI WT WT KIVI WT WT +/- +/- 2.3B KIYI WT WT KIYI WT WT ++/- 2.3B KIIA WT WT KIIA WT WT -+ 2.3B KIIV WT WT KIIV WT WT -+/- 2.3B KIIY1 WT WT KIIY WT WT ++ 2.3B 1 this construct may have spontaneous release into media, but at a rhomboid cleavage site

73 Table 6. EBA P4-P2 analysis highlights importance of P4 residue

Name N-term JxtMem P5-P2 TM C-term PfR4 PfR1 Figure EBA WT WT WT WT WT WT +-2.2A MAYY WT WT MAYY WT WT +-2.4A MIYY WT WT MIYY WT WT -+2.4A MPAY WT WT MPAY WT WT +-2.4A MPIY WT WT MPIY WT WT +-2.4A MPYA WT WT MPYA WT WT ++2.4A MPYI WT WT MPYI WT WT +-2.4A MPAA WT WT MPAA WT WT ++2.4A MPII WT WT MPII WT WT +- MIII WT WT MIII WT WT --2.4A MVYY WT WT MVYY WT WT -+2.4B MLYY WT WT MLYY WT WT ++/-2.4B MFYY WT WT MFYY WT WT ++/-2.4B MT YY WT WT MTYY WT WT +/- + 2.4B MSYY WT WT MSYY WT WT +-2.4B MRYY WT WT MRYY WT WT +-2.4B MKYY WT WT MKYY WT WT +-2.4B MDYY WT WT MDYY WT WT +-2.4B MEYY WT WT MEYY WT WT +-2.4B

Table 7. Baebl P4-P2 variants induce cleavage by PfROM1

Name N-term JxtMem P5-P2 TM C-term PfR4 PfR1 Baebl WT WT WT WT WT WT +- VLVV WT WT VLVV WT WT -- VV WT WT IPVV WT WT +/- - VL WT WT VLVV WT WT + TALI WT WT TALI WT WT ++ MLEK WT WT MLEK WT WT -- MLEK + SpiJxt WT Spitz MLEK WT WT - TALI + TgAMAJxtMem WT TgAMA TALI WT WT + SpiJxtMem WT Spitz wt WT WT + Spitz JxtMem+TM WT Spitz MLEK Spitz WT + IIYF WT WT IIYF WT WT ++ IAYF WT WT IAYF WT WT +- IPIF WT WT IPIF WT WT ++ IPAF WT WT IPAF WT WT +- IPYI WT WT IPYI WT WT ++ IPYA WT WT IPYA WT WT ++

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Exploiting the rules governing PfROM4’s substrate selectivity that I uncovered in

Chapter II, I was able to generate ‘super-substrate’ mutants that enhanced proteolysis ~5-20 fold. Our laboratory also solved high-resolution crystal structures of a model rhomboid to identify boronate as a warhead that enhanced inhibition ~100-fold. Combining these observations, I designed a ‘super-substrate’ derived peptide-boronate compound with the aim of inhibiting PfROM4. Indeed, these compounds selectively inhibited PfROM4 proteolysis with μM potency; importantly, by maintaining a ‘clashing’ residue, the activity of a panel of metazoan rhomboid enzymes was unaffected.

When I added these PfROM4 selective inhibitors to malaria cultures, they disrupted host-cell invasion specifically and iteratively, such that parasite growth ceased over 2-3 cycles. Not only does this work establish for the first time that PfROM4’s activity is necessary and targetable for parasite invasion, it also provides a strategy for designing selective inhibitors for rhomboid enzymes from other organisms.

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As resistance to first-line therapies against Plasmodium falciparum emerges in both southeast Asia (Ashley et al. 2014; Noedl et al. 2008; Dondorp et al. 2009) and Africa (Lu et al. 2017), there is a pressing need for the development of new anti-malarial agents. In particular, agents with novel mechanisms of action are coveted as many of the compounds in the drug development pipeline are derivatives of existing compounds (Wells, Hooft van

Huijsduijnen, and Van Voorhis 2015; Shanks and Möhrle 2017). Given that PfROM4 appears to be essential for blood stage parasites (O’Donnell et al. 2006; Lin et al. 2013), it is an attractive therapeutic target for such an agent.

Targeting PfROM4, however, is complicated by the fact that invasion is rapid (<1 minute) (Dvorak et al. 1975; Riglar et al. 2011), offering a limited time-window for inhibition. Moreover, it remains unclear how ‘druggable’ the membrane-immersed rhomboid family of proteases are, as they have never been therapeutically targeted (Wolf and Verhelst

2016). Existing inhibitors of rhomboid proteases are not candidates for such targeting as they suffer from poor potency, non-selectivity, and the need for preincubation (S Urban,

Lee, and Freeman 2001; Zoll et al. 2014; Vinothkumar et al. 2013; Pierrat et al. 2011).

Moreover, even more recently described rhomboid inhibitors lack activity against eukaryotic enzymes (Tichá et al. 2017). Finally, rhomboids are insensitive to most other serine protease inhibitors, partly because they attack their substrates with the opposite stereochemistry from most other serine proteases (‘si’ rather than ‘re’ face attack) (S Urban 2010).

To target PfROM4, then, a new approach is needed. Here, I combined my understanding of how PfROM4 recognizes its substrate with other innovations in our laboratory to design a selective, non-toxic inhibitor of PfROM4.

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Engineering a ‘super-substrate’ by iterative optimization

Our laboratory has recently found that mutations that enhance natural substrate processing increase the potency of substrate-based inhibitors derived from those modified sequences (Cho, Dickey, and Urban 2016). Based on this observation, I began with the P4

EBA-175 variants (Arg, Asp, Asn) from the previous chapter that enhanced processing by

PfROM4, and subjected them to additional rounds of mutagenesis. These residues were selected so as to start with diverse amino acid chemistry at this position. My aim was to iteratively optimize the substrate so as to identify the sequence that most enhanced PfROM4 processing, and use that as a template to design a potent, selective substrate-based inhibitor.

I found that the addition of a serine at the P2 position enhanced cleavage from ~3- fold for the single amino acid substitution to ~5-fold for the combined P4/P2 substrate mutants (Figure 3.1A, sequences depicted on right, quantification not shown). In confirming that these constructs were recognized by PfROM4 like the native substrate, I found that while the cleavage site for the P4-P2 sequences DYS and QYS remained intact, that for RYS shifted (Figure 3.1B). Rhomboid lacks physiological affinity for its substrates, and mutations often induce cleavage site shifts (Moin and Urban 2012). Because I was attempting to identify a binding sequence, I discarded RYS and focused my downstream analysis on the

DYS and QYS constructs. Importantly, I avoided P3 substitutions as those induced cleavage by hR2 (see previous chapter), and would therefore reduce the selectivity of an inhibitor that incorporated such a residue.

While studying Spitz processing, I found that positively charged residues in the juxtamembrane region (P10-P5) of EBA-175 enhanced PfROM4 processing (data not shown). When I incorporated lysine and arginine into these positions of the QYS and DYS constructs, I found that they further enhanced cleavage of QYS but not DYS or QYY

(Figure 3.1C, data not shown), indicating that combinatorial enhancement was dependent on the identity of both P4 and P2 residues. When I created a construct that incorporated all enhancing residues, I found that it was processed ~15-20-fold better than the wildtype EBA-

175 substrate (Figure 3.1C, blue arrow, denoted KRR QYS in sequence on right). I also confirmed that the cleavage site of this ‘super-substrate’ remained intact (Figure 3.1D), validating that this template could be used for subsequent inhibitor design.

Boronate warheads enhance potency of substrate-based inhibitors >100-fold

The next step in designing a substrate-based inhibitor was to choose the right warhead to target the enzyme’s unique active site (Walker and Lynas 2001). In addition to its alternative attack stereochemistry, rhomboid also differs from all other known serine proteases in that it stabilizes the oxyanion with a tripartite interaction. A previous attempt at designing mechanism-based inhibitors had used peptides conjugated to a chloromethylketone (CMK) group to inhibit EcGlpG (Zoll et al. 2014), but these compounds made a double covalent adduct with the enzyme and disrupted the oxyanion hole. They also required preincubation and had and an IC50 of >100 μM. Our group unexpectedly found that aldehydes (CHO) served as more natural oxyanion mimics than

CMKs (Cho, Dickey, and Urban 2016) and did not need preincubation. Even then, however, they had an IC50 of ~100 μM. In searching for more potent warheads, we turned to the boronate group because it harbored two hydroxyl groups that could potentially serve as an oxyanion and an additional hydrogen bond donor, respectively. Additionally, boronate containing compounds had previously found to be more potent inhibitors in a variety of other serine proteases, including those that attack from the ‘si’ face (Smoum et al. 2012;

Walker and Lynas 2001).

We first asked if boronates indeed exhibited the hypothetical bond-forming advantages we had hypothesized. To do so, Sang Cho in our lab co-crystallized the EcGlpG enzyme with tetrapeptides conjugated to a boronate, CMK, and CHO group (Figure 3.2A).

Although this enzyme only shares ~9% sequence identity with PfROM4, their active sites are structurally conserved and EcGlpG has served as a model enzyme for the field. For the

CMK, he was able to resolve both the single linkage to the catalytic serine and the double linkage to both catalytic residues (Figure 3.2B). While the singly-linked CMK (Figure 3.2C),

CHO (Figure 3.2D), and boronate (Figure 3.2E) all formed the tripartite oxyanion stabilizing bonds, only the boronate formed an extra hydrogen bond with the catalytic histidine (Figure

3.2E, highlighted in green). These results provided structural validation of our choice of warhead.

To confirm that these structural advantages of boronate-warheads corresponded with increased potency of inhibition, Sin Urban tested these tetrapeptide-warheads for their ability to inhibit GlpG in E. coli cells and found that the tetrapeptide-boronate was ~200- fold more potent in its EC50 compared to the tetrapeptide-CHO (Figure 3.2F, dose-response curves depicted on right), and was fully reversible (Figure 3.2G). A hexapeptide-boronate enhanced potency over the tetrapeptide-boronate by a factor of ~10 (Figure 3.2H, purple curve in dose-response curve), suggesting that extending the peptide could enhance potency, presumably through more interactions with the enzyme.

RiBn is a non-toxic, PfROM4 selective inhibitor

Combining these observations, we designed a decapeptide derived from our ‘super- substrate’ (Figure 3.1) conjugated to a boronate warhead (structure and sequence depicted in

Figure 3.3A). This compound, termed RiBn for rhomboid-inhibiting boronate, inhibited both EBA-175 and AMA-1 (a PfROM4-cleavable variant) processing by PfROM4 (Figure

3.3B) with an EC50 of ~27uM (Figure 3.3B, right). Inhibition was specific to the boronate warhead as the ‘super-substrate’ sequence linked to the aldehyde did not inhibit PfROM4

(Figure 3.3C). RiBn was non-toxic, did not alter the morphology (Figure 3.3D), or replication (Figure 3.3E) of HEK cells.

I next tested RiBn’s ability to inhibit other rhomboid enzymes, including four that are active at the cell-surface. Reassuringly, RiBn had no discernable effect on the activity of any other rhomboid enzyme tested, including the other active rhomboid enzyme in malaria,

PfROM1 (Figure 3.3F). Trafficking, glycosylation, and secretion of all substrates was also unaffected. By contrast, PfROM4’s closest homolog, TgROM5, was uninhibited by the compound. These results demonstrated that I had succeeded in my goal of designing a selective, non-toxic PfROM4 inhibitor.

RiBn is a malaria invasion inhibitor

When treated with RiBn, malaria growth was suppressed in a standard growth assay

(Figure 3.4A) with a EC50 and EC90 of 37μM and 64μM, respectively. As with inhibition of

PfROM4 activity, suppression of parasitemia was specific to the boronate warhead; the peptide-CHO had significantly reduced potency (Figure 3.4B). Additionally, the compound did not affect the morphology or viability of the erythrocytes (Figure 3.4C). While these data suggested that the effects of RiBn on parasites was due to inhibition of PfROM4 and not toxic or off-target effects, characterizing the precise susceptibility of the parasites to RiBn would provide further support that this compound was indeed acting through PfROM4 inhibition.

My expectation was that consistent with prior evidence (reviewed in (Sinisa Urban

2009; Sibley 2013)), PfROM4’s activity would be most critical during invasion of erythrocytes by blood stage parasites. Thus, parasites should be most susceptible to RiBn

81 compound during invasion. To test this directly, I added RiBn to schizont-enriched erythrocytes and monitored invasion efficiency ~24-hours later. When analyzed by flow- cytometry analysis, there was a dose-dependent reduction in invaded erythrocytes with a

~50% reduction at the highest dose tested (Figure 3.4D). Moreover, the morphology of the

RiBn-treated parasites that did invade suggested that invasion may have been incomplete:

RiBn-treated parasites that did invade appeared to be ‘stuck’ to the plasma membrane

(Figure 3.6E). This finding could be consistent with incomplete maturation of the parasitophorous vacuole if, as expected, PfROM4’s substrates accumulated on the posterior end of the vacuole without the enzyme’s activity, thereby preventing membrane fusion.

RiBn-treated parasites exhibit a delay in growth

A ~50% reduction in invasion efficiency of RiBn-treated parasites was not sufficient to explain why parasites ceased growth in the initial drug assay (Figure 3.1A). Thus, I followed RiBn-treated parasites by flow-cytometry over the course of multiple invasive cycles to look for other defects. I found that after the initial invasion defect, the RiBn-treated parasites that did successfully invade also exhibited a growth defect: at 18-hours after treatment, they were less mature than control treated parasites as assessed by [DNA] (figure

3.5A, red shading). At 42-hours, control-treated parasites were mature and ready to egress whereas those treated with RiBn still exhibited a growth defect (blue shading). The defect was most striking at the 56-hour timepoint where control-treated parasites had reinvaded while RiBn-treated parasites remained in the 1st invasive cycle (orange shading).

To determine if I could rescue this phenotype, I washed out RiBn at 18-hours post invasion. During the course of the 1st invasive cycle, RiBn-treated parasites mirrored those where the compound had been washed out (Figure 3.5B, 42-hour time point). Both remained delayed when compared to the control-treatment by flow-cytometry and imaging.

When given a chance to reinvade, parasites where RiBn had been washed out at 18-hours showed a partial rescue in both the invasion efficiency as well as growth whereas RiBn- treated parasites were reduced in number and delayed. This experiment reiterated that parasites were most sensitive to RiBn during invasion since washing out the compound did not rescue the defect until the next invasive cycle.

I confirmed this hypothesis by adding in RiBn 18-hours after parasites had already invaded. As expected, RiBn did not affect these parasites until the subsequent round of invasion (Figure 3.6A). Morphology of added in RiBn was similar to control treatment at 42 hours (Figure 3.6B). This effect was reflected in the parasitemia curve (Figure 3.6C).

RiBn does not affect egress of parasites from erythrocytes

Reinvasion consists of both egress from infected erythrocytes as well as invasion of new erythrocytes. Like invasion, egress is also dependent on proteolysis and protease inhibitors like the cysteine protease inhibitor E-64 inhibit egress (Blackman and Carruthers

2013; Boyle et al. 2010). Because my data did not disentangle egress from invasion, I could not rule out the possibility that RiBn affected egress, possibly through a non-PfROM4 dependent mechanism. Thus, I treated heparin synchronized, magnetically purified schizonts

(Boyle et al. 2010) with RiBn and compared it to E-64 (positive control) and RPMI (negative control). After 8 hours of treatment, E-64 caused invaded cells to stall at egress whereas

RiBn treatment mirrored control-treated parasites (Figure 3.6D, erythrocytes stalled at egress shown in box). By disentangling egress from invasion, I conclusively demonstrated that RiBn acts specifically during invasion, consistent with PfROM4 inhibition.

RiBn acts iteratively to cure even artemisinin-resistant cultures of malaria

Finally, to assess if targeting PfROM4 is a viable therapeutic strategy, I examined the effects of prolonged culturing of parasites in presence of RiBn. After treatment with just a

83 single-dose of RiBn, cultures were cured of malaria after 2-3 invasive cycles (Figure 3.7A, growth curves shown in 3.7B). The artemisinin-resistant strain of malaria, C580Y (Straimer et al. 2015), was also susceptible to RiBn (Figure 3.7C), demonstrating that PfROM4 activity is essential for even this critical strain.

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In this chapter, I leveraged our understanding of rhomboid substrate specificity, enzymology, and structural biology to design a non-toxic PfROM4 selective inhibitor from first principles. This chemical genetic tool was used to not only decipher the role of

PfROM4 in Plasmodium falciparum blood-stage parasites, but to also demonstrate that

PfROM4 is both essential and targetable. This represents the first therapeutic targeting of any rhomboid enzyme (Wolf and Verhelst 2016).

While PfROM4 has been thought to be essential for blood-stage parasites

(O’Donnell et al. 2006; Lin et al. 2013), genetic analysis only revealed that knockouts were unable to generated; thus, its precise function remained mostly unelucidated. By designing a

PfROM4 selective inhibitor, I was able to isolate PfROM4’s function as being critical to invasive stage parasites, consistent with our a priori hypothesis of its role in ‘capping’ proteolysis, allowing successful formation of the parasitophorous vacuole (Baker, Wijetilaka, and Urban 2006; O’Donnell et al. 2006; Figure 3.8). Importantly, chemical inhibition of

PfROM4 was sufficient clear parasitemia without the aid of antibodies targeted to PfROM4 substrates; it had been hypothesized that a primary role of PfROM4-mediated shedding of surface antigens was immune evasion (Rugarabamu et al. 2015; Shen et al. 2014).

This work also establishes the ‘druggability’ of PfROM4. It had been previously considered a poor drug target given its membrane immersed active site and narrow window of activity – invasion typically proceeds in <1 minute (Dvorak et al. 1975; Riglar et al. 2011).

The combination of a targeting sequence with a boronate-warhead likely afforded us rapid- onset kinetics, allowing for physiological, selective inhibition; prior rhomboid inhibitors were nonselective and required preincubation, limiting their utility in therapeutic targeting (Zoll et al. 2014; Wolf et al. 2015; Pierrat et al. 2011). The boron group itself has been utilized in clinically approved compounds to inhibit other hydrolases. These include the proteasomal inhibitor bortezomib (Smoum et al. 2012) and, more recently, the Ƣ-lactamase inhibitor vaborbactam (Lomovskaya et al. 2017; Hecker et al. 2015). Thus, with further medical chemistry optimization, RiBn could serve as a starting point to develop a clinical compound targeting PfROM4.

Finally, not only does this work demonstrate that PfROM4 is druggable and essential for invasion, it also lays the groundwork to take a similar chemical biology approach to target other rhomboid enzymes. While eukaryotic rhomboid enzymes are thought to play an important role in the pathogenesis of a variety of human pathogens (Sinisa Urban 2009), limited genetic and biochemical tools have precluded both an analysis of their function or therapeutic targeting. Many of these enzymes, though, are active in our heterologous cleavage assay (Baker, Wijetilaka, and Urban 2006; Baxt et al. 2008; Brossier et al. 2005;

Riestra et al. 2015), allowing for similar substrate-optimization approaches to design chemical biology tools. The approach described in this chapter, therefore, could not only define the function of a whole host of previously poorly characterized rhomboid enzymes, but could also establish a platform to target them therapeutically.

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Sin Urban designed and guided the overall project. Sin Urban and I designed the

‘super-substrate’ experiments and I performed all those experiments. Sangwoo Cho co- crystallized EcGlpG with the respective peptide-warhead compounds and performed the

85 structural analysis. Sin Urban designed and performed the in vivo GlpG assay in E. coli cells.

Sin Urban and I designed all experiments related to RiBn; I performed all those experiments.

I thank Dr. Slavica Pavlovic-Djuranovic for teaching me malaria culturing techniques and

Rosanna Baker for her help in culturing parasites while I was unable to be in lab. I also thank the laboratory of David Sullivan (JHSPH) for providing the C580Y artemisinin-resistant parasites.

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DNA constructs

Open reading frames were cloned into pcDNA3.1 (Invitrogen) as N-terminal GFP- tagged/C-terminal Flag-tagged substrates or N-terminal 3xHA-tagged rhomboid proteases.

Expression was driven by a CMV promoter. EBA-175, AMA1, PfROM1/4 and TvROM1 were all recoded for human expression as described previously (Baker, Wijetilaka, and Urban

2006). Additionally, EBA-175 and AMA1 were cloned without their large ectodomains to both help with expression and streamline the analysis. Mutants were generated either via

QuikChange Site-Directed Mutagenesis (Agilent Genomics) or an inverse PCR strategy. All constructs were confirmed by DNA sequencing (Genewiz).

Heterologous HEK-cell based cleavage assay

Cleavage of these substrates and the activity of enzymes was determined in a slightly modified heterologous cleavage assay (Baker, Wijetilaka, and Urban 2006). Briefly,

86 conditioned in 400μL of serum-free DMEM (Sigma) containing the appropriate compound where indicated. For cells transfected with AMA1, the metalloprotease inhibitor BB-94

(20μM) was included to reduce spontaneous ectodomain shedding. Media containing the

GFP-tagged N-terminal cleavage product and their respective cells were harvested ~16-20-h later by re-suspension of lyophilized media and lysis of cells in Laemmli buffer. Protein from lysates and media were resolved on 4%-12% gradient Bis-Tris gels in MES running buffer

(Life Technology), and transferred onto nitrocellulose membranes with a Trans-Blot

(BioRad) semi-dry system. After blocking (Li-COR Biosciences), membranes were probed with anti-GFP (Abcam, ab32146), anti-HA (Roche, 3F10), or anti-FLAG (Sigma, F7424) primary antibodies followed by anti-rabbit/rat secondary antibodies conjugated to infrared fluorophores (Li-COR Biosciences) and imaged on an Odyssey infrared scanner (Li-COR

Biosciences). Two-color quantitative blots are depicted.

Cleavage site analysis

To determine the cleavage site of rhomboid substrates, HEK293T cells were transfected in 6-well plates as described above, but with a higher amount of enzyme: substrate:enzyme ~50:1. 18-hours post-transfection, cells were washed 1x with DMEM conditioned with serum-free media (DMEM). ~24 hours later, cells were resuspended in

DMEM, pooled, pelleted (1000gx10minutes), and lysed in RIPA buffer in the presence of a protease inhibitor cocktail (Complete Tablets, Roche). After homogenization in a cup horn sonicator (Branson) and filtration through a loading tip, samples were subjected to anti-

FLAG immunopurification (Sigma). Briefly, 3xwashed anti-FLAG beads were incubated with samples for 1-1.5hrs at 4C. Agarose beads were washed 2x with RIPA buffer and 2x with ddH20. Bound peptide was eluted with a 4:4:1 mix of H2O:Isopropanol:Formic acid.

Eluent was spotted onto a sinapinic acid matrix, and analyzed by MALDI-TOF mass

87 spectrometry on a standards calibrated Voyager DE Instrument (AB SCIEX) as previously described (Moin and Urban 2012). Resultant spectra were analyzed and plotted in the R environment with aid of the Maldiquant package (Gibb and Strimmer 2012).

Inhibition of Endogenous GlpG in E. coli Cells

Analysis of GlpG rhomboid proteolysis in living E. coli cells was assayed as recently described (Cho, Dickey, and Urban 2016). Briefly, E. coli cells (wildtype NR698 strain and its

ƅGlpG sister strain) harboring the pBAD-TatA-Flag plasmid were grown shaking at 250rpm and 37°C in Lauria broth supplemented with ampicillin until culture optical density monitored at 600nm reached 0.4-0.5. TatA-Flag expression was induced with 25 μM L- arabinose, the indicated compounds were added directly to the cultures, and cultures were grown for an additional 2 hours shaking at 250 rpm and 37°C. Cells were lysed in reducing and denaturing TricineSDS buffer, proteins were resolved on 16% Tricine polyacrylamide gels (Invitrogen), and electrotransferred to nitrocellulose (Bio-Rad). TatA-Flag was detected with anti-Flag antibodies (Sigma, F7425) and secondary antibodies conjugated to

IRDye800cw, and quantified on an Odyssey infrared fluorescence laser scanner (Li-COR

Biosciences).

Fitting and Statistical Analysis of Dose-Response Curves

Dose-response curves were analyzed in the R environment. Curve fitting and graphing was performed with the DRC package and, unless indicated, dose-response curves were fitted with a 4-parameter logistic regression model, accounting for top and bottom baselines, the EC50 and the slope of the curve. Shaded regions reflect 95% CI of the fit.

Where indicated, points represent averages of multiple biological experiments. The EC50 is depicted along with its SE derived from the fit.

P. falciparum Culture and Invasion Assays

Asexual P. falciparum 3d7 and C580Y parasites (kind gift of Dr. David Sullivan, Johns

Hopkins School of Public Health) were cultured at 3-5% hematocrit in RPMI (with 0.5% w/v Albumax II (Gibco), hypoxanthine, gentamicin, and HEPES) under the presence of a

90%|5%|5% N2|O2|CO2 gas mixture at 37C. O-donor blood was either purchased from

Interstate Blood Bank, Inc., or obtained from the Johns Hopkins School of Public Health.

Ring-stage parasites were synchronized with 5% sorbitol once a week, or as needed for experiments.

For invasion assays, synchronized, schizont-stage parasites were magnetically purified as previously described (Bates et al., 2010) (MACS Miltenyi Biotec). Briefly, mature-stage parasites at 3-5% parasitemia were pelleted, resuspended in RPMI, and run on a pre- equilibrated LS column (Miltenyi Biotec). Bound parasitized erythrocytes were eluted with culture media and yield was calculated on a hemocytometer. Parasitized erythrocytes were then incubated at the appropriate parasitemia (0.1-0.2% for dose-response curves) in a 96- well plate in the presence or absence of the peptide-warhead inhibitors at 1.5% hematocrit.

Total sample volume was 100 μL. The plate was placed in a humidified culture chamber under presence of a 90%|5%|5% N2|O2|CO2 gas mixture at 37C.

The generate tightly synchronized parasites for the egress assay, parasites were given a 6-hour window to invade without the invasion-inhibitor heparin (200μg/mL, Sigma) present as previously described (Boyle et al. 2010). Segmented, mature parasites were then magnetically purified on CS column with a SuperMACS magnet (Miltenyi Biotec) as described above. Schizont-enriched parasites were then incubated with either 10μM E-64

(Sigma), RiBn or RPMI control at 1.0% parasitemia, 1.5% hematocrit in a 96-well plate and cultured as above. After ~8 hours, cultures were examined for the fraction of stalled, egressing parasites by flow cytometry as described below.

Flow cytometry analysis for absolute parasitemia determination and growth characterization was performed by resuspending static cultures in the 96-well plates by shaking at 275 RPM x 10 minutes in an Excella shaker (New Brunsiwick), removing a small volume of culture (typically 7.5μL), washing cells in 1xPBS, resuspending them in 1xPBS containing 0.1Ƭg/mL acridine orange and monitoring FL1/FL3 fluorescence on a

FACSCalibur instrument (BD Biosciences). Methanol-fixed, acridine orange stained parasitized erythrocytes were imaged on an inverted fluorescence microscope (Nikon) with bandpass filters. Images were overlaid in Photoshop CC (Adobe). All flow cytometry data were analyzed in FlowJo.

Analysis of RiBn Effects on Human Cells

Cell viability of RiBn-treated HEK293T cells was determined with a LIVE/DEAD

Viability/Cytotoxicity Kit (Molecular Probes). Calcein AM and ethidium homodimer-1 stained cells were imaged on an inverted fluorescence microscope (Nikon) with the appropriate bandpass filters. Flow cytometry on the same samples was performed with a

FACSCalibur instrument (BD Biosciences). For cell cycle distribution determination, RiBn

+/- treated HEK293T cells growing in serum were washed in 1xPBS, fixed in ice-cold 70% ethanol, RNase A treated and stained, with 50 Ƭg/mL propidium iodine. Flow cytometry was performed as above by monitoring the FL2/FL3 channels.

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