Investigating the Mechanism of Action of Artemisinin Antimalarials and the Role of Ferriprotoporphyrin Ix Heme

INVESTIGATING THE MECHANISM OF ACTION OF ARTEMISININ ANTIMALARIALS AND THE ROLE OF FERRIPROTOPORPHYRIN IX HEME

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

Laura Elizabeth Heller, B.A.

Washington, DC April 15, 2019

INVESTIGATING THE MECHANISM OF ACTION OF ARTEMISININ ANTIMALARIALS AND THE ROLE OF FERRIPROTOPORPHYRIN IX HEME

Laura Elizabeth Heller, B.A.

Thesis Advisor: Paul D. Roepe, Ph.D.

ABSTRACT

Malaria continues to be a serious threat to human health. The current first-line treatment for Plasmodium falciparum malaria is one of several available artemisinin (Art) combination therapies. Artemisinin, derived from the Chinese herb Artemisia annua, contains a highly reactive 1,2,4 trioxane ring. The exact mechanism for activation of this artemisinin pharmacophore and its relation to parasite death remain an active area of research. In its reduced form, free ferriprotoporphyrin heme (FPIX) is known to catalyze endoperoxide cleavage for artemisinin drugs. The activated drug likely proceeds from an oxy radical form to a carbon centered radical form that is then capable of alkylating a variety of targets within malarial parasites. It has been proposed for some time that FPIX liberated upon hemoglobin catabolism is one such target. Tragically, ACT resistance is emerging in some regions, and is typically defined clinically as a “delayed clearance phenotype” (DCP).

Using optimized extraction procedures, mass spectrometry and UV-Visible spectroscopy,

I have quantified the abundance of free FPIX and hemozoin at various stages of the P. falciparum life cycle for artemisinin-sensitive and delayed clearance phenotype parasites.

Additionally, I have identified artemisinin drug-FPIX adducts from bolus dosed parasites for the first time. My results show that along with altered intraerythrocytic development, DCP parasites show altered levels of free FPIX relative to the isogenic control throughout the intraerythrocytic cell cycle, which can lead to fewer FPIX− DHA adducts formed within live parasite. Where non-

iii crystalline FPIX is likely an important target of the Art drugs, I have also synthesized two simple

Art probes in order to better define drug localization, drug – target binding, and influx / efflux of these drugs. I also monitored the reaction between the Art derivative artesunate (ATS) and FPIX in the presence of glutathione (GSH). The reaction is observed to slow dramatically in the presence of other relevant FPIX-binding antimalarial drugs. My results show that the rate of ATS activation is limited by the rate of reduction of the FPIX Fe(III) center by GSH and other drugs can compete with GSH for association with FPIX. These data are important for defining the molecular pharmacology of Art drugs and the mechanism of evolving Art resistance.

ACKNOWLEDGEMENTS

Thank you to my mentor Dr. Paul Roepe for pushing me to be the scientist I always hoped to become. To my best friends, thank you for your empathy, laughter, and unwaivering support.

Many thanks, Laura Heller

vii TABLE OF CONTENTS

CHAPTER 1. INTRODUCTION ...... 1

1.1 General Background and Significance ...... 1

1.2 Plasmodium Biology ...... 3

1.2.1 Acquisition of Host Cell Cytoplasm ...... 4

1.2.2 Degradation of Hemoglobin via Cysteine Proteases ...... 6

1.2.3 Degradation of Hemoglobin via Aspartic Acid Proteases ...... 9

1.2.4 Hemozoin (Hz) Formation ...... 11

1.3 Antimalarial Drug Classes and Pharmacology ...... 13

1.3.1 Quinoline Antimalarials ...... 14

1.3.2 Antifolates and Antifolate Combinations ...... 16

1.3.3 Aryl Alcohols ...... 17

1.3.4 Artemisinins and Artemisinin Combination Therapies ...... 18

1.4 P. falciparum Drug Resistance ...... 20

1.4.1 Chloroquine Resistance (CQR) ...... 21

1.4.2 Other Quinoline Drug Resistances ...... 22

1.4.3 Cytostatic vs. Cytocidal Resistance ...... 23

1.4.4 Resistance to Antifolates and Aryl Alcohols ...... 24

1.4.5 Evolving Artemisinin Resistance (ArtR) ...... 25

1.5 Artemisinin Pharmacology and Mechanism of Artemisinin Resistance ...... 27

1.5.1 Heme Artemisinin Interactions and Activation of Drug ...... 27

1.5.2 Effects of Hemoglobin Metabolism and Glutathione on Artemisinin Potency ...... 32

v 1.5.3 Altered Ferriprotoporphyrin IX Heme Interactions or Hemoglobin Digestion in

Artemisinin Resistant Parasites ...... 33

1.5.4 PfK13 Mutations and PfVps34 ...... 34

1.5.5 Proteomics Studies Suggesting Multiple Art-Based Drug Targets ...... 37

1.6 Purpose of This Study ...... 40

CHAPTER 2. MATERIALS AND METHODS ...... 42

2.1 Materials ...... 42

2.2 Methods...... 43

2.2.1 Parasite Culturing ...... 43

2.2.2 Extraction and Purification of Heme and Hemozoin ...... 43

2.2.3 Live Parasite Drug Treatment ...... 45

2.2.4 Mass Spectrometry ...... 45

2.2.5 Hz Quantification by UV-Visible Spectroscopy ...... 46

2.2.6 Model Compound Studies ...... 47

2.2.7 ATS Activation NMR Studies ...... 49

2.2.8 Synthesis of Art Drug Probes ...... 50

2.2.8.1 Synthesis of Biotinylated Artesunate Probe ...... 50

2.2.8.2 Synthesis of Alkynyl Dihydroartemisinin Probe ...... 50

2.2.9 IC50 Assays to Determine Biological Activity of Probes ...... 52

2.2.10 Subcellular Fluorescence Imaging of Art Drug Probes ...... 53

2.2.10.1 Fluorescence Imaging of Biotinylated Artesunate Probe ...... 53

2.2.10.2 Fluorescence Imaging of Alkynyl Dihydroartemisinin Probe ...... 54

vi CHAPTER 3. QUANTIFICATION OF FREE FERRIPROTOPORPHYRIN IX HEME AND HEMOZOIN FOR ARTEMISININ SENSITIVE VS DELAYED CLEARANCE PHENOTYPE PLASMODIUM FALCIAPRUM MALARIAL PARASITES ...... 56

3.1 Introduction ...... 56

3.2 Results ...... 59

3.3 Discussion ...... 72

CHAPTER 4. DIHYDROARTEMISININ-FERRIPROTOPORPHYRIN IX ADDUCT ABUNDANCE IN PLASMODIUM FALCIPARUM MALARIAL PARASITES AND THE RELATIONSHIP TO EMERGING ARTEMISININ RESISTANCE ...... 76

4.1 Introduction ...... 76

4.2 Results ...... 80

4.3 Discussion ...... 94

CHAPTER 5. ARTESUNATE ACTIVATION BY HEME IN AN AQUEOUS MEDIUM ...... 101

5.1 Introduction ...... 101

5.2 Results ...... 106

5.3 Discussion ...... 115

CHAPTER 6. SYNTHESIS AND ANALYSIS OF TWO ARTEMISININ DRUG PROBES: LOCALIZATION AND COMPETITION ANALYZED VIA SPINNING DISK CONFOCAL MICROSCOPY ...... 119

6.1 Introduction ...... 119

6.2 Results ...... 122

6.3 Discussion ...... 130

CHAPTER 7. CONCLUSIONS ...... 134

APPENDIX I: PERMISSION CORRESPONDENCE ...... 142

BIBLIOGRAPHY ...... 159

vii LIST OF FIGURES

Figure 1.1. Distribution of P. falciparum around the world ...... 1

Figure 1.2. P. falciparum life cycle ...... 2

Figure 1.3. Kinetics of the P. falciparum life cycle ...... 3

Figure 1.4. Endocytosis and the cytostome in the parasitized RBC ...... 5

Figure 1.5. Hemoglobin degradation in P. falciparum and conversion to free heme ...... 11

Figure 1.6. Structure of Hemozoin solved by powder X-ray diffraction ...... 12

Figure 1.7. Structures of common quinoline antimalarial drugs ...... 14

Figure 1.8. Structures of antifolates, sulfadoxine and atovaquone ...... 16

Figure 1.9. Structures of aryl alcohol antimalarials ...... 17

Figure 1.10. Structures of artemisinin derivatives ...... 19

Figure 1.11. Spread of chloroquine resistance ...... 24

Figure 1.12. Partial map of South East Asia with an overlay of the Greater Mekong Basin highlighted in blue ...... 26

Figure 1.13. Structure of dihydroartemisinin (DHA) with numbered C3 and C4 carbons ...... 28

Figure 1.14. Reaction between ferrous iron and dihydroartemisinin to form iron-oxo species with initial oxygen radical and rearrangement to the C4 radical ...... 30

Figure 1.15. Reaction between ferrous heme (Fe(II)PPIX) and dihydroartemisinin to form a covalent adduct ...... 31

Figure 2.1. Lysis of parasites and multiple washes yields a parasite pellet ...... 44

Figure 2.2. Free ferriprotoporphyrin IX heme is separated from hemozoin after incubating and shaking with warm 4% sodium dodecyl sulfate for 2.5 hours ...... 44

Figure 2.3. UV-Visible absorbance spectra of parasite-extracted hemozoin ...... 47

v Figure 2.4 UV-Visible absorbance spectra of pure Hemin-Cl used to calibrate the absorbance of hemozoin samples versus concentration ...... 47

Figure 2.5. Monitoring the reaction of ferriprotoporphyrin IX heme and glutathione ...... 48

Scheme 2.1. One-pot synthesis of biotinylated artesunate probe ...... 50

Scheme 2.2. Synthesis of terminal alkynyl artemisinin ...... 51

Scheme 2.3. Addition of fluorophore to clickable alkynyl artemisinin ...... 52

Figure 3.1. Giemsa smears of live parasites across the life cycle ...... 60

Figure 3.2. Kinetics of intraerythrocytic development for isogenic control strain versus delayed clearance phenotype parasites ...... 61

Figure 3.3. Kinetics of intraerythrocytic development for additional control and delayed clearance phenotype strains ...... 61

Figure 3.4A. Mass spectrum of pure ferriprotoporphyrin IX heme ...... 64

Figure 3.4B. Mass spectrometry fragmentation pattern of pure ferriprotoporphyrin IX heme .....64

Figure 3.5. Mass spectrometry calibration curve for free heme using pure hemin-Cl ...... 65

Figure 3.6. Mass spectrum of pure heme solubilized from β-Hematin (synthesized in vitro) using

NaOH ...... 66

Figure 3.7A. Mass spectrum of free heme isolated from synchronized CamWT trophozoites .....67

Figure 3.7B. Mass spectrum of free heme solubilized from isolated parasite hemozoin ...... 68

Figure 3.7C. Mass spectrum of chloroform extracts of non infected red blood cells ...... 68

Figure 3.8. Free heme abundance across the parasite intraerythrocytic life cycle ...... 70

Figure 3.9. Quantifying hemozoin by UV-Visible absorbance spectroscopy ...... 71

Figure 3.10. Levels of hemozoin across the parasite intraerythrocytic life cycle ...... 71

Figure 3.11. Overlays of free heme and hemozoin production ...... 74

vi Figure 4.1. Intermediates in the production of ferriprotoporphyrin IX – dihydroartemisinin covalent adduct ...... 81

Figure 4.2. Absorbance spectra of ferriprotoporphyrin IX heme after addition of glutathione and dihydroartemisinin ...... 83

Figure 4.3. Time dependent reaction between ferriprotoporphyrin IX heme, glutathione, and dihydroartemisinin as monitored by UV-Visible absorbance spectroscopy ...... 84

Figure 4.4A. Mass spectrum of heme – dihydroartemisinin adduct synthesized in vitro ...... 85

Figure 4.4B. Mass spectrometry ionization of heme – dihydroartemisinin adduct synthesized in vitro ...... 86

Figure 4.4C. Mass spectrum of the reaction between heme and dihydroartemisinin without addition of glutathione ...... 86

Figure 4.4D. Mass spectrometry calibration curve of ferriprotoporphyrin IX – dihydroartemisinin adduct ...... 87

Figure 4.5A. Adduct isolated from parasites after bolus dosing parasites with 5 µM dihydroartemisinin for 6 hours ...... 88

vii Figure 4.7. Quantification of ferriprotoporphyrin IX – dihydroartemisinin adducts for control

CamWT and Cam3.11-R539T-Revertant strains versus delayed clearance phenotype CamWT-

C580Y and Cam.311-R539T parasites ...... 92

Figure 4.8. Mass spectrum of hemozoin isolated from CamWT parasites bolus dosed with dihydroartemisinin ...... 93

Figure 5.1. Structures of antimalarial drugs used in this analytical study ...... 101

Figure 5.2. Artesunate (ATS) and the numbering scheme employed in this work ...... 106

Figure 5.3. The 3-methyl resonance of artesunate before (a) and after (b) reaction with reduced heme formed in situ from Fe(III) heme and glutathione ...... 107

Figure 5.4. NMR resonance signal of 3-methyl of artesunate obtained in 30 minute intervals from an aqueous sample containing ferriprotoporphyrin IX heme, glutathione and artesunate ...... 108

Figure 5.5A. Conversion of artesunate to C4 centered radical artesunate versus time ...... 109

Figure 5.5B Conversion of artesunate to C4 centered radical artesunate versus time (top, ¿) and oxidation of reduced glutathione (GSH) into oxidixed glutathione (GSSG) versus time (×) .....109

Figure 5.6. Proposed mechanism for alkylation of ferriprotoporphyrin IX heme by artesunate. 110

Figure 5.7. The amount of unreacted artesunate as a function of time in the absence () vs presence of other FPIX - binding antimalarial drugs ...... 111

Figure 5.8. The amount of unreacted artesunate as a function of time in the presence of increasing ratios of chloroquine...... 112

Figure 5.9A. Proposed structures for ferriprotoporphyrin IX heme - artemether adducts formed ...... 114

Figure 5.9B. Mass spectrometry peaks observed for ferriprotoporphyrin IX heme - artemether adduct formation in the presence of Epi Quinine ...... 114

viii Figure 6.1. A drug probe derivatized with an alkynyl tag ...... 120

Figure 6.2. An alkynyl artemisinin probe ...... 121

Figure 6.3. Biotinylated probe retains its antimalarial potency as quantified by IC50 assay, compared with underivatized artesunate against P. falciparum strains HB3 and Dd2 ...... 123

Figure 6.4. Alkynyl probe retains antimalarial potency via IC50 assay, relative to artemether against P. falciparum strains HB3 and Dd2 ...... 124

Figure 6.5. Localization of ATS-b probe in P. falciparum infected red blood cells ...... 125

Figure 6.6. Localization of DHA-a probe in P. falciparum infected red blood cells ...... 126

Figure 6.7. Competition experiments between biotinylated artesunate probe versus underivatized artesunate in P. falciparum infected red blood cells ...... 128

Figure 6.8. Time-dependent localization of DHA-a probe ...... 129

Figure 6.9. Schematic Representation of a trophozoite-infected red blood cell, drawn approximately to scale ...... 132

ix LIST OF TABLES

Table 1.1. Genes up-regulated in ArtR parasites as observed by Rocamora et al...... 34

Table 1.2. Genes down-regulated in ArtR parasites as observed by Rocamora et al...... 34

Table 1.3. Overlapping protein targets of Art found by Ismail et al. and Wang et al...... 39

Table 1.4. Top hits for Up- or Down-Regulated proteins found in three studies for 3D7 parasites ...... 40

Table 5.1. Observed m/z peaks for FPIX-ATM adduct formation in the presence of EPI, detected via electrospray ionization (ESI) mass spectrometry ...... 115

Table 6.1. IC50 values for ATS and ATS-b probe ...... 123

Table 6.2. IC50 values for ATM and DHA-a probe ...... 124

x LIST OF ABBREVIATIONS

ABPP, activity-based protein profiling

AcO, acetate or acetyl group

ACT, Artemisinin combination therapy

ALLN, N-acetyl-L-leucyl-L-leucyl- L-norleucinal

AQ, Amodiaquine

Art, Artemisinin

ArtR, Artemisinin Resistance

ArtS, Art-sensitive

ATM, Artemether

ATQ, Atovaquone

ATS-b, Biotinylated ATS

ATS, Artesunate

BiPY, 2,2′-bipyridyl

CDCl3, Chloroform

CQ, Chloroquine

CQR, CQ resistance

DCP, Delayed Clearance Phenotype

DFP, Deferiprone

DHA-a, Alkynyl DHA probe

DHA, Dihydroartemisinin

DHFR, Dihydrofolate Reductase

DHPS, Dihydropteroate Synthase

xi DMAP, 4-Dimethylaminopyridine

DMSO, Dimethyl Sulfoxide

DTT, Dithiothreitol

DV, Digestive vacuole

EDC·HCl, N-(3-Dimethylaminopropyl)-N′-Ethylcarbodiimide Hydrochloride

EPI, 9-Epi-Quinine

EPR, Electron Paramagnetic Resonance

ESI-MS, Electrospray Ionization Mass Spectrometry

EtOAC, Ethyl Acetate

FPIX, Ferriprotoporphyrin IX

GGCS, Gamma Glutamylcysteine Synthetase

GSH, Glutathione

GSSG, Oxidized Glutathione Dimer

Hb, Hemoglobin

Hf, Halofantrine

HPI, Hours Post Invasion

HRP, Histidine-rich Protein

Ht, Hematocrit

HTA, Heme-targeted Antimalarial

Hz, Hemozoin

IC50, Half-maximal inhibitory concentration

LD50, Half-maximal lethal concentration

Lum, Lumefrantrine

xii MOA, Mechanism of Action

MQ, Mefloquine

MRP, Multidrug Resistance Protein

NMR, Nuclear Magnetic Resonance

P, Parasitemia

PABA, p-Aminobenzoic Acid

PC, Phosphatidyl Choline

PfCRT, Plasmodium falciparum chloroquine resistance transporter

PfPI3K, Plasmodium falciparum phosphatidyl-3-kinase

PfVPS34, Plasmodium falciparum vacuolar protein sorting 34(gene/PROTEIN)

PG, Proguanil

PI3′P, Phosphatidyl 3′-Phosphate

PPM, Parasite Plasma Membrane

PPQ, Piperaquine

PQ, Primaquine

PVM, Parasitophorous Vacuolar Membrane

PY, Pyrimethamine

PYN, Pyronaridine

QN, Quinine

QNR, Quinine Resistance

QNS, Sensitive to QN

RBC, Red Blood Cell

RPM, RBC Plasma Membrane

xiii RSA, Ring Stage Survival Assay

S.A., South America

S.E.A., South East Asia

SDCM, Spinning Disk Confocal Microscopy

SDS, Sodium Dodecyl Sulfate

SNP, Single Nucleotide Polymorphism

SX, Sulfadoxine

T1, Longitudinal Relaxation Time

TCP, Trophozoite Cysteine Protease

TMS, Tetramethylsilane

WHO, World Health Organization

xiv 1 CHAPTER 1. INTRODUCTION

1.1 General Background and Significance

One of the world’s most deadliest diseases, malaria causes over 200 million clinical cases and at least 500 thousand deaths annually. It is a vector-borne disease caused by an intracellular protozoan parasite of the genus Plasmodium and transmitted by the female Anopheles mosquito.

Of the six species of Plasmodium that cause malaria in humans (P. ovale, P. knowlesi, P. malariae, P. vivax, P. falciparum, and P. cynomolgi), P. vivax is the most widely distributed species and the cause of 25-40% of malaria cases worldwide [Gething et al., 2011]. P. falciparum infections, however, are the most lethal [Francis et al., 1997].

Figure 1.1. Distribution of P. falciparum and P. vivax around the world. Areas in red, purple, and orange represent countries affected by either P. falciparum malaria, P.vivax malaria, or both. Reproduced from [Feachem et al., 2004] with permission.

Young children without acquired malaria immunity and pregnant women are particularly susceptible to the complications associated with P. falciparum infection. Most cases are in sub-

1Sumitted in part to Tropical Medicine & Infectious Disease in April 2019 by Laura E. Heller and Paul D. Roepe*

1 Saharan Africa and parts of South East Asia (S.E.A.).

P. falciparum is transmitted when a malaria-infected female mosquito feeds on a human, transferring infectious sporozoites in its saliva to the human bloodstream (Figure 1.2).

Figure 1.2 P. falciparum life cycle. Credit CDC, Alexander J. da Silva, PhD, and Melanie Moser, 2002 (CDC, 2010). Public domain content available from the Public Health Image Library (PHIL), ID# 3405.

These sporozoites invade hepatocytes and develop in the human liver for two weeks before being released as large clusters of new merozoites that then invade and infect red blood cells (RBCs).

Once inside the red blood cell, parasites pass through a cycle of developmental stages called ring, trophozoite, and schizont. At schizogony, the red blood cell ruptures, releasing new merozoites and re-infection of new RBCs by the released merozoites occurs. This cycle lasts approximately 48 hours (Figure 1.3). While the intraerythrocytic cycle is purely asexual, these

2 parasites can also differentiate into a sexual stage called gametocytes, where male and female forms both exist (micro- and macro-gametocytes, respectively). These gametocytes can be ingested by the female anopheline mosquito during blood meal of an infected human, where they then sexually reproduce in the midgut of the mosquito. In this process oocysts form and eventually rupture to release sporozoites, which can migrate to the salivary glands of the mosquito. In a subsequent blood meal they inoculate the next human host and continue the parasite life cycle.

Figure 1.3 Kinetics of the P. falciparum life cycle. Giemsa stained parasites showing the development of parasites throughout the erythrocytic life cycle. From left to right, early ring (0 Hours Post Invasion (HPI)), late ring (12 HPI), mid trophozoite (24 HPI), early Schizont (36 HPI), and late Schizont (48 HPI).

1.2 Plasmodium Biology

Malarial parasites are adept at surviving and proliferating within the hemoglobin-rich cytoplasm of red blood cells (RBCs). Their survival depends on efficient catabolism of most all red blood cell Hemoglobin (Hb). As proposed by Coy Fitch, there are at least three main processes involved with Hb catabolism: (1) ingestion of red blood cell Hb and traffic to the parasite digestive vacuole (DV), (2) digestion of Hb and subsequent release of amino acids and free ferriprotoporphyrin IX (FPIX), and (3) the sequestration of FPIX as a crystallized pigment referred to as hemozoin (Hz) [Fitch et al., 1998]. Hemozoin is formed in the DV where the pH is

~ 5.5 [Dzekunov et al., 2000], an acidic environment in which FPIX is insoluble.

3 1.2.1 Acquisition of Host Cell Cytoplasm

Plasmodium parasites utilize erythrocyte Hb as a main source of amino acid nutrients.

Parasites are thought to degrade almost all host Hb during the ring and trophozoite stages of development. Hb physically enters the parasite through invaginations that form pinocytotic vacuoles. In 1965 using electron micrographs, Maria Rudzinska and William Trager first found evidence of host cytoplasm engulfed by invagination of the parasite plasma membrane, subsequently forming a DV in P. berghei and P. falciparum malaria [Rudzinska et al., 1965].

The process was described as pinocytosis, from the Greek word ‘pinein’ meaning 'to drink”

(contrasted with phagocytosis which stems from the word ‘phagein’, i.e. to eat). The described phenomenon was later described in more detail [Elliott et al., 2008] using higher resolution TEM images. Even from these original images however, it was clear that the DV was the site of Hb digestion and hemozoin formation. One year after observing the pinocytosis of red blood cell Hb,

Sprinz et al. published the first known observation of a structure on the surface of the parasite which they called the cytostome. A cytostome (from cyto for cell and stome for mouth) is microtubule-supported groove through which food is directed into vacuoles. The parasite cytostome was first observed on the outer surface of the parasite, spanning the double membrane of the parasite separating parasite and erythrocyte cytoplasm (Figure 1.4), and attaching to the inner parasite membrane over the entire surface of the merozoite. The cytostome grows as the parasite life cycle progresses, and holds the host cell cytoplasm within its double membrane

[Aikawa et al., 1966]. Eventually, a new DV can pinch off from the cytostome, with only a single membrane envelope [Rudzinska et al., 1965].

4 Cytostome Endosome

RBC Plasma Membrane

Parasitophorous Vacuolar Membrane

DV Parasite Plasma Nucleus Membrane

Hz Parasitophorous Vacuole

Figure. 1.4 Endocytosis and the cytostome in the parasitized red blood cell.

Degradation of host red blood cell Hb to free amino acids is important for the malarial parasite as the parasite cannot scavenge sufficient free amino acids from the blood and the parasite has only a limited capacity for the synthesis of amino acids. It is clear that the parasite degrades Hb for amino acid uptake as the Hb content of a red cell decreases significantly (up to

80%) [Ball et al., 1948] during the life cycle of the intraerythrocytic parasite. There is also an obvious spike in the concentration of free amino acids within the parasite, which has been observed qualitatively and quantitatively [Sherman et al., 1966] to be of the same type and quantity of the sums of those amino acids in the red blood cell. Rates of consumption of red blood cell oxygen and glucose increase with parasitemia, suggesting these substances are also scavenged by the parasite. In fact, parasitized red blood cells can use up to 75x more oxygen and glucose than is required by a normal red blood cell. The parasite obtains its nitrogen through the

5 breakdown of Hb as well. While there is almost no Hb breakdown in the RBCs of uninfected horses and chickens, those infected with P. gallinaceum malaria show rapid hydrolysis of Hb

[Moulder et al., 1946]. Lastly, amino acid composition in the parasite is similar to that of the host red blood cell cytoplasm, and feeding erythrocytes radiolabeled amino acids leads to their incorporation into parasite proteins [Zarchin et al., 1986].

1.2.2 Degradation of Hemoglobin via Cysteine Proteases

When parasites digest Hb in the DV, FPIX heme is converted to crystalline hemozoin

(Hz) while globin is hydrolyzed to its constituent amino acids (Figure 1.5). The DV is acidic

[Krogstad et al., 1985 and Benett et al., 2004] similar to that of mammalian lysosomes where cysteine and aspartic acid proteinases hydrolyze proteins under these acidic conditions [Bond et al., 1987]. It was first determined in 1988 by Rosenthal and colleagues that a malarial cysteine protease in necessary for degradation of Hb to FPIX heme and globin polypeptides [Rosenthal et al., 1988]. Leupeptin and E-64, two peptide inhibitors of cysteine proteases, inhibited Hb proteolysis and led to a buildup of undegraded host cytoplasm in the digestive vacuole, specifically globin monomers. This suggested that the inhibition of the protease specifically blocked degradation. Intact globin is not found in non-E-64-treated parasites [Rosenthal et al.,

1988]. Importantly, using these inhibitors to block cysteine protease activity not only blocked Hb digestion but also parasite growth, suggesting that if the parasite could not digest erythrocyte Hb then it did not have the amino acids necessary to synthesize proteins required for parasite development and replication [Rosenthal et al., 1991]. The degree of cysteine protease inhibition is also proportional, unsurprisingly, to the degree of inhibition of parasite growth [Rosenthal et al., 1989]. This phenomenon has been observed in P. falciparum and other Plasmodium species, where cysteine proteases play the same role [Rosenthal et al., 1993]. In 1995 Rosenthal and

6 colleagues identified the first cysteine protease involved in Hb degradation: falcipain-1 (formally termed TCP, for trophozoite cysteine protease) [Salas et al., 1995]. Falcipain-1 inhibition caused the DV to swell and fill with undegraded Hb. Treatment of synchronized ring-stage parasites with micromolar levels of cysteine protease inhibitors such as E-64 and leupeptin for 20 h led to an accumulation of ~2 pg of globin per parasite, which represents about 10% of red blood cell globin. Ginsburg and colleagues suggested that Hb can spontaneously denature under acidic conditions, but this is only true at pH ≤ 4.0 and not necessarily at a physiologically relevant pH range [Gabay et al., 1993]. Rosenthal and colleagues incubated Hb at 37 °C for 4 hours at pH 5.0 and above, which did not change the appearance of Hb on nondenaturing gels [Gamboa et al.,

1996]. This supported the interpretation that enzymatic activity (and not only the acidic pH of the vacuole) is responsible for the extensive Hb degradation occurring in the parasite.

Hb degradation occurs in distinct steps. It is first broken down from tetramer form into truncated alpha- and beta-globin chains bound to heme [Rosenthal et al., 1996]. After dissociation of the tetramer into monomers, heme is released. Rosenthal and colleagues found that the cysteine protease inhibitor E-64 was able to block both the dissociation of the tetramer as well as the release of heme from globin [Gamboa et al., 1996]. In 2000, Rosenthal and colleagues purified falcipain-2 from parasite DVs using affinity chromatography and suggested falcipain-2 alone was responsible for up to 90% of the cysteine protease activity in isolated trophozoites. Interestingly, the authors did not initially detect two other known falcipains during purification (now known as falcipains-1 and -3) by either sequencing or mass spectrometry.

Importantly however, falcipain-2 was more effective at cleaving denatured Hb rather than native

Hb, suggesting it may not participate in the first step of Hb catabolism. In 2001, a third putative cysteine protease (falcipain-3) was characterized and was found to be homologous to falcipains -

7 1 and -2 [Siwali et al., 2001]. Where maximum transcription of falcipains -1 and -2 occurs in early trophozoites, transcription of falcipain-3 peaks in late trophozoites and early schizonts

[Siwali et al., 2001]. Falcipains are mostly detected in the DV, with a small fraction detected in the cytosol [Shenai et al., 2000]. Falcipain-3 actively hydrolyzes Hb, but it is much less active against random peptide substrates relative to falcipain-2 [Siwali et al., 2001]. Both enzymes, however, still degrade globin much more readily than native Hb. Analysis by densitometry suggests that falcipain-2 was about 1.8 times more concentrated in trophozoites than falcipain-3

[Siwali et al., 2001]. Where falcipains -2 and -3 are about 65% identical in their catalytic domains, [Siwali et al., 2001] falcipain-1 is less similar (28% identity to falcipain-2). Not surprisingly, functionally it is very different as well. Greenbaum et al. characterized falcipain-1, finding that it is predominantly active during the merozoite and ring stages rather than the trophozoite stage [Greenbaum et al., 2002]. Falcipain-1 was also found at the apical end of the parasite and not found in the DV, according to immunofluorescence localization studies of parasites stained with an antibody to falcipain-1. Treatment of parasites with a selective falcipain-1 inhibitor, YA29-Eps(S,S) [Greenbaum et al., 2002] caused a dose-dependent decrease in the percent of new ring-stage parasites but did not block the life cycle development of surviving parasites, as parasites successfully completed schizogeny and rupture to form new merozoites. Thus it seems like falcipain-1 is capable of contributing to Hb degradation but it is not absolutely required for this process, and instead it may have a critical role in the invasion of merozoites to form new ring stage parasites. The P. falciparum DV contains several additional proteases including the aspartic acid protease plasmepsins which also participate in Hb hydrolysis. These enzymes cleave denatured Hb in vitro, and Hb hydrolysis in DV extracts is additive for cysteine and aspartic acid proteases [McKerrow et al., 1993].

8 1.2.3 Degradation of Hemoglobin via Aspartic Acid Proteases

In 1990, Goldberg and colleagues [Goldberg et al., 1990] purified the DV and measured hemoglobinase activity. The aspartic acid protease inhibitor pepstatin inhibited Hb proteolysis by

67%. Goldberg et al. suggested (in agreement with Rosenthal) that there are several steps involved in the breakdown of Hb, and specifically that there is an initial cleavage step which generates large Hb fragments due to aspartic protease activity (Figure 1.5). Pepstatin is said to block up to 80% of the initial cleavage (more effectively than cysteine protease inhibitors such as

E-64), but inhibits the subsequent generation of small fragments to a lesser extent. Additionally, under non-reducing conditions falcipain has little reactivity against native Hb but is extremely effective once Hb is denatured [Gluzman et al., 1994], where the activity of plasmepsins is unaffected by the presence or absence of reducing agents [Francis et al., 1996]. Taken together, these data suggested that an aspartic acid protease rather than a cysteine protease might be responsible for the first step in Hb degradation, with cysteine proteases involved in subsequent steps (Figure 1.5).

Continuing with this thought, Goldberg et al. isolated the first aspartic protease from P. falciparum trophozoite extracts, which when incubated with intact Hb cut solely at the 33Phe-

34Leu peptide bond at first, and after these two peptides are generated the protease cleaves additional fragments [Goldberg et al., 1991]. This highly specific cut at the 33-34 peptide bond is in the Hb hinge region. When Hb binds oxygen, one alpha-beta dimer swivels about 15 degrees around the other dimer, disrupting intermolecular bonding nearby with the exception of this hinge region, so that the tetramer stays intact. The scission of the 33Phe-34Leu peptide bond thus leads to an unraveling of the molecule and subsequent exposure of additional sites for proteolysis, but Hb initially stays in tetramer form. Importantly, hydrolysis of Hb by the isolated

9 protease is halted in the presence of pepstatin but not by other classes of protease inhibitors, which have been seen to be more effective only after intact Hb has been partially degraded

[Goldberg et al., 1990] (Figure 1.5). The purified protease also shares NH2-terminal homology with known aspartic proteases, where 9 of 22 residues are identical to renin, a mammalian aspartic protease [Goldberg et al., 1991]. No beta-chain cleavage was detected with this purified protease, suggesting that there is likely at least a second protease involved in the initial step of

Hb digestion [Goldberg et al., 1991].

Where it had been shown previously that aspartic acid protease inhibitors can inhibit the majority of Hb degradation (60-80%), Goldberg and colleagues [Gluzman et al., 1994] found that the cysteine protease inhibitor E-64 in combination with aspartic hemoglobinase inhibitors can block all detectable proteolysis. Inhibition of plasmepsin I by a peptidomimetic inhibitor SC-

50083 led to parasite death in culture, suggesting that plasmepsin I is essential for parasite development [Francis et al., 1994]. Additionally, IC50 assays were performed using SC-50083 and pepstatin (a general aspartic protease inhibitor) as well as E-64, and isobolograms showed synergy between cysteine and aspartic protease inhibitors where the IC50 of E-64 was lowered substantially in the presence of SC-50083. Purification of plasmepsins I and II by Goldberg and colleagues suggested that they each cleave the 33Phe-34Leu peptide bond, after which plasmepsin I cleaves the 105-106 bond and plasmepsin II cleaves at 108-109. Similar to the falcipains which are metabolically active during different stages of the parasite life cycle, the plasmepsins are expressed and processed during different stages. According to northern blot analysis of plasmepsin mRNA, plasmepsin I mRNA is most abundant during the ring stage and can be detected as early as 3 hours post red blood cell invasion. For plasmepsin II however, mRNA is not detected until 12 hours post invasion and is most abundant during the trophozoite

10 stage. Synthesis of both plasmepsins I and II peaks in the trophozoite stage and ends by the schizont stage.

N N Plasmepsins I & II Fe N N

Polypeptides O OH O OH Plasmepsins I, II, & IV Hemoglobin Heme Falcipains 1, 2, & 3 Falcilysin DPAP1

Amino Acids

Hemozoin Figure 1.5. Hemoglobin degradation in P. falciparum and conversion to free heme. Reproduced from [Gorka et al., 2013c] with permission.

1.2.4 Hemozoin (Hz) Formation

During the intraerythrocytic stages, P. falciparum digests up to 80% of host Hb. This process releases amino acids and also an oxidized form of ferriprotoporphyrin IX (FPIX) heme which is toxic to biological membranes [Tappel et al., 1953]. FPIX can initiate a chain reaction by scission of fatty acid peroxides into two free radicals [Tappel et al., 1953]. However, parasites are able to digest Hb with impunity by sequestering oxidized FPIX to form hemozoin (Hz), an inert non-toxic crystalline dimer of FPIX heme [Pagola et al., 2000]. In 1987 Fitch and

Kanjanananggulpan [Fitch et al., 1987] were the first to determine that hemozoin consisted of pure FPIX and not protein. They performed elemental analysis, determining that it is actually composed of oxygenated heme, as purified hemozoin contained about 88% FPIX and 12% O2.

11 Hemozoin is insoluble in aqueous sodium bicarbonate, but can be solubilized by sodium hydroxide [Fitch et al., 1987]. Fitch suggested that hemozoin and a material known as β-hematin were very similar [Fitch et al., 1987], though he could not prove they were identical. In 1991,

Slater et al. tested the structure of hemozoin, noting that it was identical to that of β-hematin by powder X-ray diffraction [Slater et al., 1991]. Isolating parasite hemozoin by centrifugation, incubating with 2% SDS to separate any free FPIX from the pellet, and then treating with proteinases to digest any contaminants, Slater isolated pure hemozoin. IR analysis suggested that hemozoin consisted of FPIX monomers linked via carboxylate side chain oxygens of one FPIX and the iron center of another, making an insoluble polymer. Slater et al. described hemozoin as a polymer where linkage was as they described, i.e. between propionate groups and iron centers.

However, in 2000 Pagola et al. [Pagola et al., 2000] determined that hemozoin was in fact not a polymer, but rather a crystal of unique “head to tail” FPIX dimers. In this structure, one propionate of each FPIX molecule coordinates to the iron center of an adjacent FPIX molecule, and the second propionate coordinates to the propionic acid side chain of FPIX in another dimer via hydrogen bonding (Figure 1.6).

N N O Fe O O O N N

N N O Fe N N OH O O O N N Fe O O O N N

N N O OH O O Fe N N O N N Fe O O O N N

N N O Fe O O O N N

Figure 1.6. Structure of hemozoin solved via powder X-ray diffraction. Reproduced from [Pagola et al., 2000] with permission.

12 This model is consistent with the fact that hemozoin crystals have well-defined faces, which likely would not be observed for a polymer. Hemozoin crystals can be described as having a lath-like shape, i.e. a long narrow strip, where the smallest face of the crystal is always the fastest growing one. Dorn et al. noted that both lipids extracted from trophozoite extracts, as well as pre-formed synthetic β-hematin can promote hemozoin crystal growth. Interestingly, the rate of hemozoin formation is significantly faster in the presence of parasite lipid extracts [Dorn et al.,

1998]. Later studies identified hemozoin formed within neutral lipid nanospheres [Pisciotta et al., 2007] inside the DV. While lipids are involved as catalysts in the formation of hemozoin, what initiates this process remains unclear. Histidine-rich proteins (HRPs) promote the formation of hemozoin [Sullivan et al., 1996]. But genetic ablation of the gene encoding HRP is not lethal

[Dixon et al., 2011], suggesting that there may be other proteins involved. Initially Sullivan and colleagues proposed that HRPs are secreted by the parasite into the red blood cell cytosol and then brought into the DV during the Hb digestion process. When Hb is degraded by the parasite,

HRPs might then bind to free FPIX and initiate the formation of hemozoin. Conversely, while other proteins such as bovine serum albumin can bind to FPIX, they do not initiate FPIX crystallization [Sullivan et al., 1996].

1.3 Antimalarial Drug Classes and Pharmacology

Malaria (or a disease resembling it) has been noted for over 4000 years, with the first symptoms of malaria being described in ancient Chinese writings as early as 2700 BC. However the first formal treatment for malaria was not known for several centuries. In the early 1600s, ground up bark from the cinchona tree was put into use to abate fevers due to malaria. By 1820, the small molecule quinine (QN) had been isolated and purified from the tree bark and was used as an effective treatment against malaria [Kyle et al., 1974]. QN remained the front-line

13 antimalarial until World War II when it was largely replaced by chloroquine (CQ; Figure 1.7).

Similar in structure but with a less narrow therapeutic range and lower toxicity, this new quinoline was developed by German scientists as a replacement to QN in 1934 [Gregson et al.,

2005] and by U.S. scientists at the National Institutes of Health shortly after [Slater et al., 2006].

Additional drugs based on the QN structure were subsequently discovered (Figure 1.7) [Gregson et al., 2005].

N N HO N HO N HN HN O O

Cl N Cl N N N Chloroquine Amodiaquine Quinine Quinidine (CQ) (AQ) (QN) (QD)

HN N N HO O N N H N N N HN N CF3 NH2 CF3 Cl Cl Mefloquine Primaquine Piperaquine (MQ) (PQ) (PPQ)

Figure 1.7 Structures of common quinoline antimalarial drugs.

1.3.1 Quinoline Antimalarials

Quinoline antimalarial drugs act on the blood stages of P. falciparum, specifically the stages that actively degrade Hb. [Zhang et al., 1986] Quinolines are weakly basic (pKa’s for CQ are 10.23 and 8.15), allowing both neutral and charged forms to be present in the parasite. In their neutral form quinolines such as CQ can passively diffuse through membranes, and upon entering the DV

(pH 5.2-5.6) become preferentially charged and less permeant [Ursos et al., 2002; Krogstad et al., 1986]. Quinoline-based antimalarial drugs (Figure 1.7) hyperconcentrate in the digestive vacuole of the parasite [Yayon et al., 1985; Ursos et al., 2002] and bind to free FPIX within the

14 DV after its release from digested red blood cell Hb [Gorka et al., 2013a; et al., Casabianca et al., 2004; Alumasa et al., 2011; de Dios et al., 2003]. Free FPIX is toxic [Loria et al., 1999; Fitch et al., 1983], but as discussed the malarial parasite sequesters FPIX as nontoxic hemozoin. The

“ferriprotoporphyrin interaction hypothesis”, suggests that CQ binds to free FPIX, inhibiting hemozoin crystallization and leading to an accumulation of free FPIX which is intrinsically toxic to the parasite [Macomber et al., 1967; Fitch et al., 1986].

It was initially proposed [Schueler et al., 1964] that formation of a CQ-FPIX complex would detoxify CQ. However the opposite is true, as it was determined [Macomber et al., 1967] that the binding of CQ to FPIX is what gives CQ its antimalarial activity at nanomolar concentrations. Formation of a dative CQ-FPIX complex likely contributes to accumulation of free FPIX by making it unavailable to serve as substrate for the growing hemozoin crystal.

[Slater et al., 1992; de Dios et al., 2003]. Slater and Cerami measured hemozoin crystal growth, using trophozoite extracts as the catalyst [Slater et al., 1992]. In the presence of quinoline antimalarials such as CQ, they were able to completely inhibit hemozoin formation. The specific mechanism by which CQ or other quinolines prevent FPIX crystallization is not certain, though it is thought to bind to free FPIX [Leed et al., 2000; de Dios et al., 2003] and to also directly interact with the growing face of the hemozoin crystal [Sullivan et al., 1996b]. Radioactive CQ, when incubated in parasites, is found almost exclusively over the hemozoin crystals within the

DV [Sullivan et al., 1996b]. If quinolines prefer to interact with the growing hemozoin chain in such a way, it is likely that the drugs can incorporate into the growing crystal itself, blocking further addition of FPIX units and leaving toxic free heme within the parasite DV. Interestingly, in CQR strains where CQ accumulation into the DV is less pronounced, [Paguio et al., 2009] CQ

15 association with hemozoin is less pronounced [Sullivan et al., 1996b] suggesting that something about the resistance phenotype limits drug association with the growing hemozoin crystal.

Several 4-aminoquinoline derivatives have been shown to localize to the parasite DV of living parasites [Aikawa et al., 1972; Cabrera et al., 2009] and these quinolines’ affinity and specificity for binding to FPIX has been found to agree well with their affinity and specificity for binding to iRBCs [Fitch 1969, Fitch 1970; Chou et al., 1980] It is generally accepted that the mechanism of action of quinolines such as QN, MQ and AQ is similar to that of CQ, where they each interfere with hemozoin crystallization, though their interaction with heme may not be the only source of potent cytotoxic activity. [Warhurst et al., 2007] Quinoline methanols such as QN and QD show similar potencies for inhibition of β-hematin formation in vitro [Gorka et al.,

2013a]. MQ is a less potent inhibitor and is thought to possibly possess a second or alternative mechanism of action, though this is yet to be determined. Unlike most quinolines, the 8- aminoquinoline primaquine (PQ) does not inhibit hemozoin formation [Dorn et al., 1998].

Instead, PQ acts against the liver stage of the parasite life cycle [Labbe et al., 2001].

1.3.2 Antifolates and Antifolate Combinations

NH 2 Cl O H O N N O N O S O H2N Cl N N NH NH

N N N OH H H H O NH2 Cl Sulfadoxine (SX) Pyrimethamine (PY) Proguanil (PG) Atovaquone (ATQ)

Figure 1.8. Structures of antifolates, sulfadoxine and atovaquone

Antifolate antimalarials such as proguanil (PG) and pyrimethamine (PY) were first developed in the late 1940s and early 1950s [Archibald 1951]. In general, these drugs work by inhibiting enzymes of the folate pathway, which results in decreased serine, methionine and

16 DNA synthesis. PG and PY inhibit dihydrofolate reductase (DHFR) [Olliaro et al., 2001]. After first introduction, antifolates were noted to be immediately effective even against CQR parasites

[Powell et al., 1963]. In the 1960s, Sulfadoxine (SX) showed promise as an antimalarial, where single 1 gram doses of the drug were effective, though fairly slow, at killing intraerythrocytic parasites [Gregson et al., 2005]. SX works by mimicking p-aminobenzoic acid (PABA) and preventing the formation of dihydropteroate [Olliaro et al., 2001]. It was soon determined that the combination of sulfadoxine and pyrimethamine was more effective than either drug alone

[Hurly 1959], i.e. the combination was synergistic. Additionally, this combination of drugs led to faster cytocidal activity than seen with sulfadoxine alone [Richards 1966 and Harinasuta et al.,

1967].

Proguanil (PG) is used in combination today with Atovaquone (ATQ), a napthalene- based antimalarial who’s main target is complex III (ubiquinol: cytochrome c oxidoreductase) which feeds the electron transport chain present in the parasite mitochondrion [Painter et al.,

2007]. The combination of ATQ + PG, known as Malarone, is currently prescribed by the CDC as a single tablet taken orally for the prevention of malaria.

1.3.3 Aryl Alcohols

F N N F OH F N OH

OH N Cl Cl HN N O

Cl Cl Cl N Cl Halofantrine (HF) Lumefantrine (LUM) Pyronaridine (PYN)

Figure 1.9. Structures of aryl alcohol antimalarials

17 Another class of antimalarials is the aryl alcohol drugs, which include Halofantrine (HF),

Lumefrantrine (LUM), and Pyronaridine (PYN). HF was developed by the U.S. Army in the

1970s [Gregson et al., 2005] and has a structure vaguely related to that of CQ (Figures 1.9 and

1.7). It is also thought to interfere with the FPIX detoxification pathway. It is effective against

CQR malaria, though its high cost and associated cardiotoxicity have limited its use. A related compound LUM has no cardiotoxicity and is thus being used currently, including in combination with artemether (ATM) to treat CQR malaria. PYN is a related acridine aryl alcohol, which is similarly being used currently in combination with artesunate (ATS). The Aryl Alcohols PYN and LUM, similar to HF, have been suggested to inhibit hemozoin formation in the malarial parasite [Ward et al., 1998; Auparakkitanon et al., 2006].

1.3.4 Artemisinins and Artemisinin Combination Therapies

Artemisinin (Art) is a sesquiterpene lactone isolated from the Artemisia annua plant, an herb employed in traditional Chinese medicine to remedy fevers [Klayman et al., 1985]. Several different derivatives and metabolites of this drug are currently in use in different settings, with these modifications performed mainly to improve the solubility of the drug in aqueous solutions

(Figure 1.9).

18 H H H H O O O O O O O O O O O O H H H H O O O O O O O OH O OH O Artemisinin Dihydroartemisinin Artemether Artesunate (ART) (DHA) (ATM) (ATS)

H Ozonide Derivatives O O H O H O O N O O NH2 O O O N O N O O S O O Artemisone OZ277 (Arterolane) OZ439 (Artefenomel)

Figure 1.10. Structures of artemisinin derivatives

Artemisinins are particularly unique in their ability to both kill the parasite and inhibit their growth by interfering with metabolic processes. This class of drug is effective against all different stages of parasites across the intraerythrocytic life cycle, with increased potency against the initial ring stage [ter Kuile et al., 1993]. Art-treated rings can also enter a state of quiescence or dormancy rather than being killed right away, and can resume growth up to weeks after drug treatment [Witkowski et al., 2010]. In addition, this class of antimalarials has the ability to impede the development of gametocytes unlike most other antimalarial drugs [Price et al., 1996].

The exact mechanism for activation of the Art pharmacophore and its relation to parasite death remain an active area of research. One theory, however, involves the cleavage of Art’s endoperoxide bridge after reacting with FPIX iron to generate free radicals which can in turn alkylate a variety of targets, resulting in the death of the parasite [Meshnick et al., 1993].

Currently, Art combination therapies or ACTs, which have been used since the mid

2000s, are the front-line therapy for treatment of P. falciparum malaria as recommended by the

19 World Health Organization (WHO). Artemisinins in general have relatively short half-lives

(around 1 hour), and in ACTs are paired with an additional drug that has a significantly longer half-life. In this way, the “parent” Art basal drug can reduce parasite burden by orders of magnitude within a few hours or days, and the second “partner” drug can reduce the likelihood of resistance and prevent recrudescence by killing the few parasites that remain [White et al.,

2004]. In the early 1990s, ATS (Figure 1.10) was combined with MQ as an effective ACT, reducing rates of malaria infection significantly along the Thai-Myanmar border [Carrara et al.,

2009]. The success of ATS-MQ led to the deployment of other combinations such as ATM-LUM and DHA-PPQ (Figures 1.8, 1.9 and 1.10).

1.4 P. falciparum Drug Resistance

Drug resistance in malaria can be defined by the parasite’s ability to “survive and/or multiply despite proper administration and adsorption of an antimalarial drug in the dose normally recommended” [WHO 1973]. Drug resistance can in theory arise from a variety of possible factors including an increase in the parasite’s ability to degrade the drug, altered expression of a drug target, or mutations in this target which then disrupt the interactions between the drug and the target, drug transport, or drug induced pathways to cell death. Drug selection pressure plays a major role in developing drug resistance, where parasites that survive after exposure to drugs pass on fortuitous resistance-conferring mutations to the next generation of parasites. To varying degrees, drug resistance has been reported for virtually every known antimalarial. Antimalarial drug resistance is often determined by blood tests, where patients given a drug treatment are monitored for parasite clearance and/or recrudescence vs. drug

[Rieckmann et al., 1989]. Resistance can also be quantified in vitro by laboratory determined

20 IC50, determining the dose as which 50% of parasite growth is inhibited [Bennett et al., 2004]. It can also be determined by DNA sequencing, which reveals genetic mutations responsible for resistance, for example mutations in the PfCRT gene that determines CQ resistance (CQR).

1.4.1 Chloroquine Resistance (CQR)

Chloroquine (CQ) has been the most extensively used antimalarial since the 1940s, though the emergence of chloroquine-resistant (CQR) malarial strains has reduced its effectiveness. CQR first developed independently in three to four areas in Southeast Asia

(S.E.A)., Oceania, and South America (S.A.) in the late 1950s and early 1960s. This was in part due to population-based dosing regimens implemented in an effort to eliminate malaria across the globe [Foley et al., 1998]. As described, CQ and other quinolines interfere with hemozoin crystallization, allowing toxic FPIX to accumulate in parasitized erythrocytes [Fitch et al., 1998].

Consequent buildup of FPIX and drug-FPIX complexes has been suggested to be associated with parasite death [Fitch 2004]. However, it is not known if FPIX accumulates to lethal levels, and drug–heme interactions are likely not the only way in which the drugs exert toxic effects

[Gaviria et al., 2013; Roepe et al., 2014]. In fact, as detailed below (Section 1.4.3) it is now understood that CQ has multiple effects and that there are multiple layers to the molecular mechanism of CQR. Resistance to the cytostatic effects of CQ is thought to be associated with reduced CQ accumulation by the parasite. Mutations in the DV membrane protein P. falciparum chloroquine resistance transporter (PfCRT) are believed to mediate the transport of CQ from the

DV. [Fidock et al., 2000; Roepe et al., 2011] However, levels of CQ found in the DV of CQR parasites do not necessarily correlate with resistance to the parasite-kill (cytocidal) effects of CQ

[Cabrera et al., 2009]. In addition, rates of CQ transport by different isoforms of mutant PfCRT do not correlate with the CQ IC50 of the strains expressing these PfCRTs [Baro et al., 2013].

21 Taken together, these data suggest that there may be additional mechanisms responsible for influencing the susceptibility of P. falciparum to CQ in addition to efflux of CQ from the parasite DV [Gaviria et al., 2013].

1.4.2 Other Quinoline Drug Resistances

Additional 4-quinolinemethanols such as amodiaquine (AQ) and mefloquine (MQ) as well as QN are still effective against many CQR parasites [Schlagenhauf et al., 2010]. AQ and

MQ are also used as ACT partner drugs, currently recommended by WHO. This suggests that the mechanism of CQR is not identical to that for resistance to other quinolines. Resistance to other quinolines has been noted in certain areas of the globe. Quinine Resistance (QNR) is not as distinct from the QNS phenotype with regards to measured differences in IC50. QNR has been linked to mutations in several genes including PfCRT, PfMDR1, and a P. falciparum sodium proton exchanger, PfNHE-1) [Ferdig et al., 2004]. Overexpression and single nucleotide polymorphisms (SNPs) in PfMDR1 have also been associated with QNR [Sidhu et al., 2005;

Sidhu et al., 2006; Reed et al., 2000]. QNR SNPs have also been noted on PfCRT [Ferdig et al.,

2004], and overexpression of PfNHE-1 has been suggested to increase cytosolic pH in QNR parasites [Bennett et al., 2007]. The PfMDR1 gene has been linked to resistance to several different quinoline drugs. Amplification of PfMDR1 is associated with MQR [Price et al., 2004], and increased copy number of PfMDR1 serves as the best overall predictor of treatment failure with MQ. PfMDR1 and PfCRT mutations have also been suggested to confer AQR [Duraisingh et al., 1997; Sa et al., 2009]. Recently, resistance to PPQ (in combination with DHA as

DHA+PPQ) has been noted and has been linked to increased Plasmepsin II/III copy number, though this is an area of current study and much more is currently being discovered about PPQR

[Bopp et al., 2018].

22 1.4.3 Cytostatic vs. Cytocidal Resistance

In the laboratory, the potencies of quinolines are studied with respect to two mechanisms

– cytostasis (inhibition of parasite growth) and cytocidality (parasite-kill) [Roepe 2014]. Where

IC50 describes the 50% growth inhibitory drug concentration, antimalarial drug potency can also be described by its cytocidal (cell kill) activity, quantified by the 50% lethal dose (LD50). As opposed to the cytostatic assay which utilizes continuous growth for 48 h in the constant presence of low concentrations of drug, in the LD50 assay the drug is delivered in bolus fashion, meaning as a relatively short pulse of drug [Gligorijevic et al., 2008]. The quinoline cytocidal assay typically involves a 6-hour bolus dose with high concentrations of drug, after which the drug is washed away and growth is measured after 48 hours without drug. In this way, where

IC50 measures relative cell growth in the presence of drug, LD50 measures cell survival.

Cytostatic and cytocidal targets for quinolines likely differ [Gaviria et al., 2013], so molecular mechanisms of resistance to cytostatic vs. cytocidal drug effects appear to differ as well [Cabrera et al., 2009; Gorka et al., 2013a; Gaviria et al., 2013]. In a study by Paguio et al., malarial strain 7G8 was seen to be sensitive to QN (QNS) with respect to IC50 but resistant with respect to LD50, suggesting different intracellular targets for each effect [Paguio et al., 2011].

Additionally, potent IC50 does not necessarily indicate potent LD50 for quinolines antimalarials

[Gorka et al., 2013b]. LD50 does not appear to have a direct correlation with hemozoin inhibition potency, though interestingly IC50 was correlated with ability to inhibit hemozoin

[Gorka et al., 2013b]. While it is largely thought that cytostatic potential is driven by the ability to inhibit hemozoin formation, this is not necessarily true of cytocidal potency. For example,

LD50 of CQ is 5-fold greater than IC50 for the CQ- sensitive strain HB3 and almost 70-fold greater than IC50 for CQ-resistant strain Dd2, which could suggest altogether different processes

23 at work [Mott et al., 2015]. Although much evidence points to CQ – FPIX interactions being crucial for cytostatic potency, mechanisms of quinoline cidality may not involve drug-FPIX interactions [Gaviria et al., 2013]. Furthermore, for synergistic or antagonistic combinations of quinolinal antimalarials, hemozoin inhibition potency may not predict antiplasmodial IC50 or

LD50 potencies [Gorka et al., 2013c]. Autophagy can be a response to starvation or oxidative stress in the parasite whereby cellular material is eventually digested within the DV or other organelles. Plasmodium falciparum parasites possess the machinery necessary for autophagy and this pathway is triggered in response to cytocidal concentrations of drugs [Gaviria et al., 2013].

Figure 1.11. Spread of chloroquine resistance. CQR spread throughout South Africa, Africa, and South East Asia. Reproduced from [Hartl et al., 2004] with permission.

1.4.4 Resistance to Antifolates and Aryl Alcohols

Though antifolates such as Pyrimethamine (PY) and Proguanil (PG) were at one point effective monotherapies, resistance to each drug has emerged. PG was used as a monotherapy in the late 1940s and early 1950s for plantation workers across South East Asia, which led to drug pressure on the parasite and subsequent PGR. PYR also emerged in South East Asia in the

24 1960s. These antifolates are DHFR inhibitors as mentioned above, while sulfa drugs are dihydropteroate synthase (DHPS) inhibitors. Certain point mutations in the DHFR coding region are correlated with resistance to antifolates. The mutation S108N in the DHFR active site has been shown to confer PYR [Peterson et al., 1997]. This mutation also leads to decreased susceptibility to cycloguanil, the active metabolite of PG, as do mutations at codons 16 and 164 of the DHFR active site [Parzy et al., 1997]. Similarly, mutations in DHPS are linked to SX resistance [Wang et al., 1997; Anderson et al., 2011]. By 1980 SP began to fail in South East

Asia and South America [Gregson et al., 2005], and resistance to SP eventually evolved worldwide due to mutations in both DHFR and DHPS genes. ATQ, which is initially used as a single agent before being combined with PG, led to drug resistance by around 1995 due to mutations in the catalytic domain of the cytochrome bc1 complex in the parasites’ mitochondrial membrane [Vaida et al., 2000]. Resistance to aryl alcohols such as halofrantrine also arose in the

1990s and was associated with the amplification of PfMDR1, as was seen with several quinolines [Nateghpour et al., 1993].

1.4.5 Evolving Artemisinin Resistance (ArtR)

Artemisinin combination therapies (ACTs) were introduced in response to multiple independent drug failures including to CQ, antifolates and mefloquine and by 2006 they were the recommended treatment for P. falciparum malaria worldwide. Unfortunately, evolving Art resistance (ArtR) has recently been found in western Cambodia and has spread to the Greater

Mekong Subregion, a transnational area of the Mekong River basin in Southeast Asia. In 2002, the failure rate after 28 days for ATS-MQ was 14%, and just two years later in 2004, the failure rate after 42 days was 21% [Denis et al., 2006a]. ATS-MQ failure was also seen in Trat province, on the Thai side of the Thai-Cambodia border. Initially, resistance to MQ alone was

25 presumed to be the problem. However, low cure rates were also being observed for ATM-LUM in nearby Battambang Province in Cambodia [Denis et al., 2006b]. To look more specifically at the problem, a study was performed where ATS was clinically administered as a monotherapy and parasite clearance was closely monitored. Despite acceptable drug levels (i.e. no issues with counterfeit drugs), parasite clearance rates were twice as slow in Cambodia than on the Thai-

Myanmar border. This delay in parasite clearance has been found to be common across mainland and Southeast Asia [Ashley et al., 2014] (Figure 1.12).

Figure 1.12. Partial map of South East Asia with an overlay of the Greater Mekong Basin highlighted in blue. Map taken with permission from Google Maps.

26 Because of some slow parasite clearance noted after ACT treatment in clinical studies, a series of genotyping studies were done to search for a genetic basis for ArtR (also known as delayed clearance phenotype or DCP). It soon became clear that ArtR is heritable [Amaratunga et al., 2012], and that it is associated with multiple founder populations [Miotto et al., 2013].

Additionally, GWAS studies initially detected a region on P. falciparum chromosome 13 that was strongly associated with delayed clearance [Cheeseman et al., 2012]. Subsequently, parasites cultured under intermittent Art pressure for five years were sequenced [Ariey et al.,

2014] and a mutation within a gene on chromosome 13 was found. This gene encodes a protein with a Kelch motif, now called PfKelch13 or PfK13. A large number of Cambodian isolates were also found to contain mutations in PfK13. It seems that the lowered effectiveness associated with PfK13mutations causes a greater number of parasites to survive initial ACT drug doses, which over time can lead to partner drug resistance and overall decline in ACT efficacy

[Woodrow et al., 2017].

1.5 Artemisinin Pharmacology and Mechanism of Artemisinin Resistance

1.5.1 Heme Artemisinin Interactions and Activation of Drug

Emerging ArtR (DCP) has focused attention on Art pharmacology and the molecular mechanism of ArtR. Before the emergence of ArtR, in 1991 Meshnick and colleagues treated parasites with [14C] Artemisinin, running the parasite lysate on an SDS page gel and found that all of the radioactivity migrated to the solvent front of the gel, suggesting that Art had bound to a target in the parasite and that the molecular mass of the bound complex was < 3000 Da, similar to what was found when incubating Art with FPIX in vitro. Using UV-Visible spectroscopy they also noted that the peak for FPIX absorbance decreases substantially in the presence of DHA.

27 They lastly noted that a solution of Art pre-incubated with FPIX lowers the antimalarial activity of Art, while FPIX alone had no activity [Meshnick et al., 1991]. The following year, Posner et al. monitored the breakdown of a simpler but comparable endoperoxide (containing only two rings compared to Art’s three) which they synthesized and labeled with oxygen-18. Oxidation was seen to lead to the breaking of the endoperoxide bridge. Furthermore, addition of iron (II) yielded an oxygen radical, which they monitored via oxygen-18. A 1-5 shift was noted to place the radical on carbon 4, “C4”, which is then alkylated. This was characterized by 13-C and 1-H

NMR and represents the first published work on the C4 radical [Posner et al., 1992].

H 4 3 O O O H O

DHA

Figure 1.13. Structure of dihydroartemisinin (DHA) with numbered C3 and C4 carbons.

That same year, Meshnick et al. used cyclic voltammetry to suggest that Art and DHA are irreversibly reduced by incubation with FPIX chloride [Zhang et al., 1992]. Meshnick also measured EPR (electron paramagnetic resonance) for Art after a 30 minute incubation with iron

(ammonium iron (II) sulfate); the EPR suggested formation of radicals, where no such signal was seen for Art alone or Fe alone [Meshnick et al., 1993]. Via SDS-PAGE autoradiograms,

Meshnick suggested that DHA (using [3H] DHA) binds covalently to serum. In the presence of

FPIX, the percent of bound drug increased by 35%, and it decreased by 10% upon addition of iron chelator DFO [Yang et al.,1993]. Looking for what else could bind covalently to the drug,

28 in 1994 the Meshnick group claimed that [14C] Art bound proteins in vitro [Yang et al., 1994].

Measuring percent radioactivity bound to purified protein, they found that 5 - 18% of added drug bound to hemoproteins such as catalase, cytochrome c, and Hb Hemoglobin [Yang et al., 1994].

Meshnick and colleagues went a step further looking for evidence of a FPIX-drug covalent adduct. Mixing FPIX chloride with Art in MeOH and incubating in the dark for 24h, they found what appeared to be FPIX-Art adduct peaks by mass spectrometry [Hong et al., 1994].

Additionally, incubation of [14C] Art with pre-isolated parasite hemozoin also led to a loss of radioactivity in the supernatant, suggesting that hemozoin binds with Art leading to an uptake of

[14C] [Hong et al., 1994]. Similarly, incubation of iRBCs with [3H] Arteether, the ether derivative of Art, showed almost no radioactivity in the supernatant but up to 85% of radioactivity was seen after washing the pellet with 1% SDS. Meshnick suggested the [3H]

Arteether bound to FPIX was released from the pellet with SDS. To test the hypothesis that a C4 radical is formed in activation of Art, the Meshnick and Posner groups together synthesized Art derivatives with one and two methyl groups on C4. This would prevent a radical from forming on C4. The methyl-substituted derivatives had no activity against parasites, suggesting that the

C4 radical scheme was in fact necessary for the drug’s activity [Posner et al., 1994].

Bernard Meunier and colleagues [Robert et al., 1997] proposed a scheme for radical formation by forming an adduct between Art and manganese(II) meso-tetraphenylporphyrin, as the comparable manganese porphyrin is more easily demetallated than iron. The presumed porphyrin-Art adduct was studied via UV-Vis and NMR, and Meunier suggested that first a O- centered radical is formed, and upon C3-C4 cleavage, a C4 radical is formed. (Figures 1.13 and

1.14) Because the radical is now on the end of an alkyl chain, it can easily alkylate many targets.

Adducts were seen using artemether (ATM) and an additional endoperoxide, BO7 [Robert et al.,

29 1998]. This scheme was also supported by Wu et al. separately [Wu et al., 1999].

3+ Fe3+ Fe 4 4 4 O endoperoxide O H C 3 O 2 O bridge breaks 3 C3-C4 cleavage Fe2+ O O O O O O 3 H3C O OH OH O OH Oxy C Radical DHA Radical 4 Complex Complex Figure 1.14. Reaction between ferrous iron and dihydroartemisinin to form iron-oxo species with initial oxygen radical and rearrangement to the C4 radical. Reproduced from [Robert et al., 2002b] with permission.

In 2001, Meunier et al. monitored the reaction between reduced heme dimethyl ester and Art in dichloromethane [Robert et al., 2001]. While this reaction was not physiologically relevant, conditions were chosen as Art is much more soluble in organic solvent as is the dimethyl ester.

Using NMR Meunier saw what he suggested to be the formation of the oxy radical, C4 radical formation, and then Art alkylation of the FPIX porphyrin. The following year Meunier et al. incubated FPIX with equimolar Art and excess glutathione (GSH) for one hour in DMSO, and reported the formation of FPIX-Art adducts with structures for each major m/z peak. Following this observation, they made similar adducts and demetallated the porphyrin in order to more closely examine the system by NMR, which would otherwise be difficult to study due to the paramagnetic iron. Here Meunier et al. outlined the first highly detailed scheme for formation of radicals using explicit chemical structures. They also performed this experiment with a series of other non-Art endoperoxide compounds, suggesting that adducts can form between these porphyrins and trioxane drugs and that the endoperoxide bridge is a key part of their pharmacological activity. One group criticized Meunier’s use of DMSO as it is not as physiologically relevant as aqueous conditions and claimed that Art derivatives with bulky tails have lower antimalarial potency, suggesting that these tails would block access to the endoperoxide bridge and leave them inactive [Haynes et al., 2005]. Meunier et al. went on to

30 synthesize several Art derivatives with bulkier tails and reported that they do, in fact, have antimalarial activity similar to ART, ATS, ATM and others [Laurent et al., 2005].

N N Fe N N

N O OH N O O OH Fe2+ O O N N O O O OH O O O OH O OH OH Oxy Radical Fe(II)-PPIX DHA Complex

N N Fe N N N N Fe O OH O OH N N HO O H3C O H2C O OH O O OH O OH O H3C O C Radical Covalent O OH 4 Adduct Complex Figure 1.15. Reaction between ferrous heme (Fe(II)PPIX) and dihydroartemisinin to form a covalent adduct. Reproduced from [Robert et al., 2002c] with permission.

Meunier and colleagues [Robert et al., 2005] presented the first direct evidence for FPIX-

Art adducts from extracts of spleens of mice infected with P. vinckei and subsequently treated with Art. Several adduct MS peaks were identified and structures inferred [Robert et al., 2005].

These correlated well to previous MS data for in vitro adducts made in their own lab as well as by Meshnick and colleagues [Hong et al., 1994]. Together, these data suggested that heme could play an important role in the mechanism of Art action.

31 1.5.2 Effects of Hemoglobin Metabolism and Glutathione on Artemisinin Potency

Since the mid 2000s, ACTs have been the front-line therapy recommended for the treatment of P. falciparum infections by the World Health Organization (WHO). Unfortunately, by around 2006 ACTs began to show some reduced efficacy as described above. As it is known that Art-based drugs are highly reactive in the presence of FPIX and that parasites digest host cell Hemoglobin releasing free FPIX, some have begun to look for connections between loss of

Art potency and altered Hemoglobin metabolism, levels of glutathione in the parasite, or other differences which could be associated altered levels of reduced heme within the parasite. The

Tilley group [Klonis et al. 2011] studied the relationship between Hemoglobin metabolism and

Art potency, suggesting that treatment with Art inhibits endocytosis of host cell cytoplasm by measuring fluorescein-dextran uptake of parasites with increasing Art (4 hour pulse). This is not found in the absence of red blood cell lysate, presumably as Art is not activated. Testing the theory that Hemoglobin digestion could be required for activity of Arts, Tilley et al. dosed 3D7 parasites with Arts in the presence of the cysteine protease inhibitor N-acetyl-L-leucyl-L-leucyl-

L-norleucinal (ALLN). Addition of ALLN appeared to minimize the effect of Arts, with up to

90% parasite survival even in the presence of 1 mM Art. In 3D7 mutants with deletions of falcipain 2, Art is less potent; after a four hour bolus dose of 800 nM Art, 20% of the initial parasitemia remains, compared to 3D7 where there are closer to 2-5% parasites remaining. These data agree with the theory that Hemoglobin metabolism aids Art potency.

The Tilley group also tested the effects of iron chelators on Art potency. [Xie et al.,

2016] Dosing 3D7 parasites with BiPY (an Fe2+ specific chelator, 2,2′-bipyridyl) or DFP

+3 (Deferiprone, specific for Fe chelation) in an LD50 assay, these iron chelators have mild but antagonistic effects on Art and DHA. They also looked at the influence of the cysteine protease

32 inhibitor, E64d, on Art potency and note that pre-incubation of E64d for one hour appeared to inhibit Art potency, and that interestingly this antagonism was most pronounced in the early ring stage. Tilley and colleagues identify falcipains 1, 2, and 3 as active in the ring stage.

In addition to Hemoglobin metabolism, Art’s reactivity with FPIX depends upon levels of glutathione within the parasite. Glutathione is synthesized de novo by gamma glutamylcysteine synthetase (ggcs). P. berghei parasites resistant to CQ or MQ show an increase in P. berghei ggcs [Perez-rosado et al., 2002] To further study the relationship between glutathione levels and Art potency, these investigators disrupted or overexpressed P. berghei ggcs in parasites [Vega-Rodriguez et al., 2015]. Pbggcs mRNA levels for overexpressed ggcs parasites were ~5 times higher than that of wild type parasites. Level of glutamylcysteine synthetase did not effect parasite growth for ED50 levels of drug, as either increased or decreased levels of ggcs did not change the ED50 for CQ or Art when compared to ED50 values for wild type parasites. Instead, a difference was seen in parasite recrudescence. After dosing mice with

20 m/kg DHA per day for four days, mice infected with WT parasites showed recrudescence in

2-3 days after treatment, as did parasites with overexpressed P. berghei ggcs. However, mice infected with the parasites with disrupted ggcs expression did not show recrudescence and parasites failed to recover. These results suggest that glutamylcysteine synthetase levels and by association glutathione levels might influence parasite response to Art treatment.

1.5.3 Altered Ferriprotoporphyrin IX Heme interactions or Hemoglobin Digestion in Artemisinin

Resistant Parasites

A metabolomics study [Mok et al., 2011] studied differences in the metabolome of ArtS vs ArtR field isolates. They found markers for down-regulation of glutathione metabolism and

Hemoglobin digestion in ArtR trophozoites, and up-regulation of Hemoglobin digestion in ArtR

33 Schizonts compared to ArtS isolates. While Mok et al. used field isolates, a more recent paper by the same group [Rocamora et al., 2018] derived two ArtR lines in the lab by drug pressuring

3D7 early rings with 900 nM Art for four hours every other life cycle. The two lines were then subject to genomic characterization to study differences between ArtS and ArtR surviving parasites. Falcipain 1 was observed to be up-regulated in ArtR parasites compared to ArtS across the parasite life cycle. Below are tables of genes specifically down- or up-regulated in the mid to late ring stage of ArtR compared to ArtS parasites [Rocamora et al., 2018].

Table 1.1. Genes Up-regulated in ArtR parasites as observed by Rocamora et al. Gene ID Up-Regulated Gene Description FDR PF3D7_0512200 glutathione synthetase 0.20 PF3D7_1224600 cytochrome c heme lyase, putative 0.23 PF3D7_0825600 cytochrome c oxidase assembly protein, putative 0.19 PF3D7_1311700 cytochrome c2 precursor, putative 0.21

Table 1.2. Genes Down-regulated in ArtR parasites as observed by Rocamora et al. Gene ID Down-Regulated Gene Description FDR PF3D7_0727200 cysteine desulfurase, putative 0.19 PF3D7_1438900 thioredoxin peroxidase 1 0.24 PF3D7_1457200 thioredoxin 1 0.12 PF3D7_1455400 hemolysin, putative 0.22 PF3D7_1419800 glutathione reductase 0.16 PF3D7_1012300 ubiquitinol-cytochrome c reductase complex 0.19 PF3D7_1352500 thioredoxin-related protein, putative 0.24 PF3D7_1011900 heme oxygenase 0.24 PF3D7_1458000 cysteine proteinase falcipain 1 0.16

1.5.4 PfK13 Mutations and PfVps34

As described previously (section 1.4.5), in 2014 a molecular marker for Art DCP was isolated and found to coincide with mutations in the propeller region of what is called the PfK13 protein

[Ariey et al., 2014 and Ashley 2014]. Traditionally, drug resistance is quantified by comparing

IC50 values, i.e. the concentration of drug required to inhibit parasite growth by 50%. Resistance to Arts is particularly difficult to measure, however, as parasites that are less susceptible to Arts

34 do not display an altered IC50 compared to sensitive parasites. As this assay cannot distinguish the ArtR phenotype, Witkowski and colleagues developed a new assay known as the RSA or ring stage assay [Witkowski et al., 2013; Straimer et al., 2015]. In this RSA, early ring-stage parasites are bolus-dosed with 700 nM DHA for 6 hours. The parasites are then washed, cultured in drug-free media, and relative survival is determined by Giemsa smears 66 hours later. The percent survival can suggest whether the parasites are resistant or susceptible to Art treatment since sensitive parasite strains have an RSA survival rate of ~0-1% while over 10% of resistant parasites survive in this assay. This assay has been used to suggest that introducing PfK13 point mutations into an ArtS CQR strain increases ring stage survival upon Art exposure [Straimer et al., 2015]. Interestingly, introducing the same K13 propeller mutation into different genetic backgrounds results in variable levels of DCP suggesting that while PfK13 mutations are necessary to explain the RSA and clinical phenotype of Art resistance, these mutations are not sufficient to account for the full observed phenotype [Straimer et al., 2015]. While it is widely accepted that PfK13 mutations are a marker of Art resistance, Straimer and colleagues suggest that there are other molecular mechanisms that add to PfK13 effects. As PfK13 has no known orthologs in other eukaryotes, its function is not clear.

A study by the Haldar group [Mbengue et al., 2015] suggested that Arts inhibit the P. falciparum phosphatidyl-3-kinase (PfPI3K). The study found that PfK13 mutant parasites appeared to have increased levels of PI3K (a modest ~1.5 to 2-fold difference). This was found to be the case for both clinical isolates and parasites with a chromosomally inserted PfK13

C580Y mutation linked to ArtR. A linear correlation was found between levels of PI3P and ArtR as measured by the ring stage survival assay (RSA), which suggested that PI3K may be directly linked to decreased sensitivity to Art compounds in mutant PfK13 parasite strains. Mbengue et

35 al. rationalizes the increase in PI3P associated with increased ArtR by suggesting that the C580Y mutation diminishes PfK13 binding to PI3K as well as ubiquitinylation-dependent degradation of

PI3K, both increasing levels of intact, unbound kinase. As validation for this theory, effects of

PfK13 on PI3P levels were further explored [Bhattacharjee et al., 2018]. Where parasites with a synthetically modified PfK13 C580Y mutation are suggested to induce ArtR, these mutations were now suggested to expand ER-PI3P vesicles, propagating them through the parasite and exporting them into the red blood cell. The authors claim [Bhattacharjee et al., 2018] that PI3P vesicles are amplified in very early rings (0-3 HPI) in PfK13 mutant parasites.

Based on these two studies, it is unclear if Art’s interaction with PfPI3K affects the activation of Art, presumably by a metal such as iron. When the binding site between PfPI3K and DHA was modeled using molecular dynamics simulations, Mbengue et al. proposed a binding model reliant on hydrogen bonding to an intact endoperoxide. This may not be physiologically relevant as the endoperoxide bridge of Art is commonly thought to break upon contact with a target. (see Section 1.5.1 and also Berman et al., 1997] While Mbengue et al. suggested that DHA binds directly to PfPI3K, they proposed that breakage of the endoperoxide bridge is not necessary for drug binding, and instead the that the intact peroxide bond is necessary for PfPI3K inhibition via hydrogen bonding between the enzyme and the drug. This theory, however, is rather unusual and seems to contradict the work of other groups [Wang et al.,

2015]. In 2017, Hassett et al. studied the binding of PfPI3K to Art targets [Hassett et al. 2017b].

PfPI3K most closely resembles Vps34, the sole class III PI3K, [Vaid et al., 2010 and Tawk et al., 2010] though PfPI3K (called PfVPS34) lacks pleckstrin homology domains and Ras binding sites that are characteristic of class I and II PI3Ks [Hassett et al., 2018]. Hassett et al. purified

PfVps34 expressed in yeast to measure PI3P production from PI [Hassett et al., 2017a] and

36 quantified Vps34 activity and drug inhibition by Arts and related ozonide compounds [Hassett et al., 2017b]. Importantly, Art had no effect on Vps34 activity in systems lacking a source of catalytic ferrous iron (either in the form of FPIX heme or FeSO4). In the presence of these iron sources, however, DHA and artemisone inhibited Vps34 activity at nM concentrations, suggesting that activation of the endoperoxide bridge by iron is required for Arts to bind

PfVps34. Interestingly, EC50 values for PfVps34 inhibition did not correlate well with

2 antiplasmodial IC50 values for Art drugs (R = 0.54) suggesting that Vps34 is not the only target of Art drugs.

PfVPS34 has also been suggested to play a role in autophagy or autophagy signaling

[Gaviria et al., 2013 and Hain et al., 2013]. Bolus doses of ATM can trigger the autophagy pathway [Mott et al., 2015]. A recent genome-wide association study identified that a locus on chr10 containing Atg18 was associated with decreased sensitivity to DHA and ATM. [Wang et al., 2016] Additionally, PI3K inhibitors have been suggested to be potent antimalarial compounds that inhibit the autophagy pathway and are synergistic with Art compounds.

[Siriwardana et al., 2016] Lastly, PfVps34 may be a particularly key component of the P. falciparum autophagic cascade, as TOR kinase activity is typically a key regulator of autophagy but no TOR homologue has been identified in the P. falciparum genome. [Gaviria 2013 et al.,

2013 and Brennand et al., 2011]

1.5.5 Proteomics Studies Suggesting Art-Based Multiple Targets

Between 2015 and 2016, two separate groups published proteomics experiments using synthesized derivatives of Art as well as a trioxane derivative, all of which could be conjugated to biotin using click chemistry, and subsequently identified targets covalently bound to Art and related endoperoxides using high resolution mass spectrometry [Wang et al., 2015; Ismail et al.,

37 2016a; Ismail et al., 2016b]. For Art probes, Wang et al., 2015 synthesized an alkynyl derivative of Art which they clicked to a biotin azide, while Ismail et al., 2016a synthesized a similar molecule as well as an azide derivative of Art which could be clicked to an alkynyl biotin. Wang et al. found 124 potential protein targets to which their synthetic Art analog bound; Ismail et al. found 49 protein targets, and 62 targets for their non-Art trioxane probe [Wang et al., 2015;

Ismail et al., 2016a; Ismail et al., 2016b]. Comparing the results of these three papers, there are

19 proteins alkylated in all three studies, listed below along with their putative locations within the P. falciparum parasite (Table 1.3). Notable protein targets covalently bound to all three probes include plasmepsins I and II and multidrug resistance protein (MRP). As seen in table 1.4, ornithine amino transferase, alpha tubulin 1 and tubulin beta chain were each significantly down- regulated in the ring stage, and the ring-infected erythrocyte surface antigen and merozoite surface protein 1 were significantly down regulated in the trophozoite stage. If these proteins are down regulated but still able to be targeted by the drug, perhaps there is a greater possibility that they are true drug targets and not simply a random target in high abundance in the parasite.

Additional proteins that are mildly up- or down regulated in the ring or trophozoite stages are shown in Table 1.4 and include actin 1, L-lactate dehydrogenase, and plasmepsins I. Of these eight total proteins, the final five (merozoite surface protein I, tubulin beta chain, actin I, L- lactate dehydrogenase and plasmepsin I) are essential. Thus, these proteins are (1) found in all three proteomics studies, (2) down-regulated to some degree, and essential. The only protein which meets all these criterion and is also localized to the DV is plasmepsin I. Interestingly, plasmepsin I would be expected to be highly associated with heme, connecting the proteomics results with the heme-Art adducts hypothesis.

Table 1.3. Overlapping protein targets of Art found by Ismail et al. and Wang et al. Putative Gene ID Proteins Found in All 3 Papers Location Dense Granules PF3D7_0102200 Ring-infected erythrocyte surface antigen (Merozoites) PF3D7_0322900 40S ribosomal protein S3A, putative Ribosome Digestive PF3D7_0523000 Multidrug resistance protein Vacuole PF3D7_0608800 Ornithine aminotransferase Cytoplasm PF3D7_0624000 Hexokinase Cytosol PF3D7_0708400 Heat shock protein 90 Cytoplasm PF3D7_0818900 Heat shock protein 70 Nucleus PF3D7_0903700 Alpha tubulin 1 Microtubule Plasma PF3D7_0930300 Merozoite surface protein 1 Membrane PF3D7_1008700 Tubulin beta chain Microtubule PF3D7_1012400 Hypoxanthine-guanine phosphoribosyltransferase Cytosol Digestive PF3D7_1015900 Enolase Vacuole Actin Filament PF3D7_1246200 Actin I / Cytoskeleton Digestive PF3D7_1311900 Vacuolar ATP synthase subunit a Vacuole PF3D7_1324900 L-lactate dehydrogenase Cytosol PF3D7_1357100 Elongation factor 1-alpha Cytoplasm Digestive PF3D7_1407900 Plasmepsin I Vacuole Digestive PF3D7_1408000 Plasmepsin II Vacuole PF3D7_1444800 Fructose-bisphosphate aldolase Cytoplasm

Table 1.4. Top Hits for Up- or Down-Regulated proteins found in all three studies for 3D7 parasites. Log 2 expression value is described, where a value of 0 suggests normal expression, and positive or negative values denote up- or down-regulation, respectively. Proteins with values more than +/- 0.75 are shown in blue, and those with values more than +/- 1.25 are shown in red. Trophozoite Ring (9 HPI) (28 HPI) Proteins Found Putative Expression Gene ID Expression Essential? in All 3 Papers Location Value (log2 Value (log2 Ratio) in 3D7 Ratio) in 3D7 Ring-infected Dense PF3D7_ erythrocyte Granules 0.8 -2.52 Unknown 0102200 surface antigen (Merozoites) PF3D7_ Ornithine Cytoplasm -1.74 0.7 No 0608800 aminotransferase PF3D7_ Alpha tubulin 1 Microtubule -1.3 -0.08 No 0903700 PF3D7_ Merozoite Plasma -0.59 -1.52 Yes 0930300 surface protein 1 Membrane PF3D7_ Tubulin beta Microtubule -1.53 -0.02 Yes 1008700 chain Actin PF3D7_ Actin I Filament / -0.55 -1.05 Yes 1246200 Cytoskeleton PF3D7_ L-lactate Cytosol -0.76 0.49 Yes 1324900 dehydrogenase PF3D7_ Digestive Plasmepsin I 1.17 -1.01 Yes 1407900 Vacuole

1.6 Purpose of This Study

Artemisinin (Art) and its derivatives are leading antimalarial drugs and yet their mechanisms of action (MOA) are poorly understood. Because of the emergence of ArtR, a clearer understanding of Art’s MOA is essential. Recent studies have shown that Art does not target just one protein but instead likely has numerous targets. In addition to the alkylation of intracellular proteins, many studies have suggested that FPIX heme itself could be an important alkylation target in addition to its role as an activator. Many successful antimalarials have targeted or disrupted this

Hemoglobin digestion pathway and it has been suggested that Arts do as well. Therefore I

40 propose that FPIX acts as both an activator and target of Arts. While Hemoglobin digestion occurs primarily after the ring stage and thus appreciable FPIX is presumably unavailable during this stage, there is less available FPIX for drug activation in the ring stage which is consistent with the fact that Art is less effective against the parasite in its ring stage. In an attempt to contribute to our expanding understanding for Art’s MOA, I have investigated the interaction between FPIX and Art derivatives, both in the parasite and through in vitro UV-visible spectroscopy, liquid chromatography mass spectrometry, NMR spectroscopy, and fluorescence microscopy. The specific interaction between Art and FPIX as well as the effects of additional quinoline drugs on this reaction was monitored by UV-visible and NMR spectroscopies. I have quantified the abundance of free FPIX and Hemozoin at various stages of the P. falciparum life cycle for ArtS vs ArtR parasites. Additionally, Artemisinin drug-heme adducts from bolus dosed parasites have been identified and quantified across the life cycle. Lastly, clickable and biotinylated Art derivatives were synthesized in order to study the binding of Art to additional non-FPIX targets. Collectively, I hope these investigations will provide valuable insight about

Art and its mechanism of action as well as the mechanism of evolving Art resistance.

41 CHAPTER 2. MATERIALS AND METHODS

2.1 Materials

Routine chemicals, media, and solvents were reagent grade or better, purchased from

Sigma-Aldrich (St. Louis, MO) or Fisher Scientific (Newark, DE), and used without further purification, unless otherwise noted. Sterile tissue culture plastics were purchased from Fisher

Scientific (Newark, DE). Hemin-Cl was purchased from Sigma-Aldrich and artemisinin (Art) drugs were purchased from Tokyo Chemical Industry (Tokyo, Japan). Red blood cells and matched human serum for parasite culture were from Valley Biomedical (Winchester, VA).

Plasmodium falciparum clone HB3 (Honduras, Chloroquine Sensitive [CQS]), Dd2 (Indochina,

Chloroquine Resistant [CQR]) were obtained from the Malaria Research and Reference Reagent

Resource Center (Manassass, VA). CamWT, CamWT-C580Y, Cam3.II-Revertant, and Cam3.II-

R539T strains of P. falciparum [Straimer et al., 2015] were a kind gift of Dr. David Fidock

(Department of Microbiology and Immunology, Columbia University).

For synthesis of drug probes, (Trimethylsilyl)-acetylene was purchased from Matrix

Scientific (Columbia, SC). Boron trifluoride (BF3) and magnesium sulfate (MgSO4) were purchased from EMD Millipore (Gibbstown, NJ). Alexa Fluor 488 Azide was purchased from

Thermo Fisher Scientific (Waltham, MA). Bathophenanthroline disulfonic acid, Poly-lysine, copper(II) sulfate, and dithiothreitol (DTT), as well as organic solvents such as dichloromethane, ethyl acetate and hexanes were all purchased from Sigma-Aldrich (St. Louis, MO). Finally, for microscopy experiments, 10X PBS, 10X PBS-Tween 20, and 100% Triton X-100 were obtained through Fisher Scientific (Pittsburgh, PA). Paraformaldehyde, 10% glutaraldehyde, and

Fluoregel mounting media with TES buffer were obtained from Electron Microscopy Sciences

(Hatfield, PA).

42 2.2 Methods

2.2.1 Parasite Culturing

P. falciparum strains were maintained as described [Trager et al., 1973] with minor modifications [Bennett et al., 2004; Gligorijevic et al., 2006; Gligorijevic et al., 2008]. Briefly, cultures were maintained under a 5% CO2 / 5% O2 / 90% N2 atmosphere at 2% hematocrit (Ht) and 1-10% parasitemia (P) in RPMI 1640 supplemented with 10% type O+ human serum, 25 mM

HEPES (pH 7.4), 23 mM NaHCO3, 11 mM glucose, 0.75 mm hypoxanthine, and 20 ug/L gentamicin. Parasitemia was monitored by Giemsa staining and adjusted every 48 hours by adding fresh RBCs. Before addition to culture RBCs were washed with incomplete media

(RPMI 1640, 24 mM NaHCO3, 11 mM glucose, 0.75 mM hypoxanthine, pH 7.4). All cultures were routinely synchronized three times per cycle by 5% D-sorbitol treatment to obtain rings as described. [Gligorijevic et al., 2006a] Parasite cell cycle populations vs time were quantified by

Giemsa smearing the synchronized cultures in triplicate every two hours over the course of 48 hours and counting the parasitized RBCs (Figure 1.3). Results shown are the average of two independent countings for each of three independently grown cultures. Differences in the kinetics of intraerythrocytic development were extracted from Gaussian curve fitting of the raw data.

2.2.2 Extraction and Purification of Heme and Hemozoin

Purification of heme from live parasite culture was essentially as described previously

[Combrinck et al., 2013; Gabryszewski et al., 2016] but with some modifications. Briefly, at different time points across the parasite life cycle, 20 mL of synchronized culture at 8.75 % parasitemia and 2% Ht was centrifuged for 5 min at 1800 rpm and parasites were released by exposing the parasite pellets to ice cold 0.1% saponin for 10 min. Samples were centrifuged

43 again to remove the majority of red blood cell lysate and pellets were then washed 3x with ice cold PBS to yield purified parasite pellets containing free FPIX and crystallized hemozoin.

Pellets were incubated with 150 µL PBS, 50 µL H2O, and 50 µL 1M HEPES, pH 7.3 for 20 min and then washed to remove any final traces of Hemoglobin (Figure 2.1).

Figure 2.1. Lysis of parasites and multiple washes yields a parasite pellet. Initial lysis gives a dark brown parasite pellet and a red supernatant containing Hemoglobin. Multiple washes of the pellet with ice cold PBS followed by incubation with a PBS/H2O/HEPES mixture yields a parasite pellet containing no visual traces of Hemoglobin.

Warm SDS (400 µL at 4% 0.1 M NaHCO3, pH 9.1) was added to the pellet and the mixture was incubated and shaken at 37 °C for 2.5 hr. This was then centrifuged at 13,000 rpm in an

Eppendorf microfuge to separate fractions containing FPIX heme and hemozoin (Figure 2.2). In order to ensure a pure final hemozoin pellet, the pellet was washed 3x with bicarbonate buffer as crystalline hemozoin will not redissolve in bicarbonate buffer and this was stored at -4oC.

Figure 2.2. Free ferriprotoporphyrin IX heme is separated from hemozoin after incubating and shaking with warm 4% sodium dodecyl sulfate for 2.5 hours.

44 The supernatant (free FPIX) was transferred to a fresh tube where 5% guanidinium-Cl was added to precipitate SDS. The mixture was centrifuged, supernatant removed, and the pellet extracted with 1.5 mL of 2:1 CHCl3:H2O. The organic layer was saved, dried en vacuo, and extracted FPIX was re - dissolved in LCMS-grade MeOH / 0.1 % formic acid for LCMS

Analysis. Hemozoin was prepared for UV-Vis analysis by dissolving in 500 µL 0.1 M NaOH, and sample was diluted 1:1000 in LCMS-grade MeOH / 0.1 % formic acid for LCMS Analysis.

2.2.3 Live Parasite Drug Treatment

In aqueous solution artemisinin (Art) and its clinically more useful derivatives artemether (ATM) and artesunate (ATS) may spontaneously decay to dihydroartemisinin (DHA). Art may also decay to deoxy-Art [Lee, et al., 1989; White 2008]. In patients, DHA exhibits a relatively short half life that can range from 1 to several hr [Newton et al., 2002; Hien et al., 2004; Geditz et al.,

2014]. To model clinical exposure of control vs DCP parasites to Art based drugs, parasites were bolus dosed (typically for 6 hr) with plasma levels of DHA (5 µM) [Newton et al., 2002; and

Khanh et al., 1999] and then lysed. Mass populations of drug-treated infected red blood cell

(iRBC) (2% Hct and 8.75% P) were then fractionated to isolate parasites as described above.

Extraction and purification of free FPIX heme in drug adduct form was as described above.

2.2.4 Mass Spectrometry

Data were obtained in the positive ion mode on an Agilent 500-MS LCMS Ion Trap mass spectrometer. The electrospray ionization source was operated at 5 kV with N2 as the nebulizing gas. First-order ESI mass spectra were recorded in the mass range m/z 400-1000. FPIX and hemozoin samples extracted from parasites were prepared for LCMS in the following manner: free FPIX extracts prepared as above were dissolved in 500 µL LCMS Grade MeOH /

0.1% formic acid. Hemozoin pellets were resuspended in 500 µL of 0.1 M NaOH and incubated

45 overnight at 37 °C to solubilize hemozoin to FPIX and then diluted 1000x into MeOH prior to

LCMS. Solutions were introduced into the mass spectrometer by a syringe pump (Hamilton), employing a 1 mL syringe at a constant flow rate of 20 µL/min. The capillary voltage and radio frequency potential were set to 92 V and 111% respectively, and the source temperature was maintained at 325 °C. Data were acquired using MS Work Station software. Calibration was done with standard solutions by using the ratio of known analyte concentration to peak areas for those samples (see Results). Standard curves for quantification of drug - FPIX adducts were generated in the following manner: standard solutions of FPIX-DHA adduct synthesized as descrbed in results were made by incubating FPIX, glutathione, and DHA in a 1:10:1 molar ratio overnight in 40% DMSO. The ratio of known adduct concentration and adduct MS peak areas

(major peak m/z 881) were plotted as described for underivitized FPIX (see Results, Chapter 4).

2.2.5 Hemozoin Quantification by UV-Visible Spectroscopy

Parasite extracted hemozoin was washed 3x with 0.1 M NaHCO3 buffer, pH 9.1 and re- dissolved in 500 µL 0.1 M NaOH by incubation at 37 °C. Samples were analyzed via UV-

Visible spectroscopy by measuring absorbance in a 1 um quartz cuvette. Raw data was baseline corrected (Figure 2.3) in order to quantify levels of hemozoin for each sample. These spectra were then compared to a calibration curve of pure Hemin-Cl in 0.1 M NaOH (Figure 2.4).

Figure 2.3. UV-Visible absorbance spectra of parasite-extracted hemozoin. Left: Raw absorbance data; Right: Baseline-corrected data. For quantification purposes, area under the baseline-corrected curve is analyzed.

Figure 2.4. UV-Visible absorbance spectra of pure hemin-Cl used to calibrate the absorbance of hemozoin samples versus concentration. Inset: Calibration curve of peak area vs. concentration of pure hemin-Cl.

2.2.6 Model Compound Studies

To probe DHA alkylation of FPIX heme via UV - vis spectroscopy, FPIX as hemin-Cl

(200 µM) was dissolved in acetate buffer (250 µM) / 40% DMSO. Glutathione (GSH) was added

47 to a final concentration of 1 mM and absorption spectra were acquired immediately. The cuvette was capped to prevent the entry of additional oxygen. UV spectra (200-1100 cm-1) were acquired every 5 min until heme was fully reduced, i.e. when the clear Fe3+ heme peak had completely disappeared and was replaced by the Fe2+ heme spectrum (see Results and Figure 2.5).

Figure 2.5. Monitoring the reaction of ferriprotoporphyrin IX heme and glutathione. Addition of glutathione leads to the reduction of Fe(III) heme at 389 nm to Fe(II) heme at 417 nm. Once reduction of FPIX was complete, artesunate was added to the reaction mixture. If left alone, as shown, FPIX would not re-oxidize under these conditions.

ATS or DHA in pure DMSO were then added (to a final 1:1 molar ratio of drug:heme), and absorbance was immediately measured. The cuvette was again capped and absorbance was measured every 5 min for the first 90 min, and then every 30 min for 12 hr more. In order to monitor production of the final adduct (FPIX-Fe(III)-C4-DHA; see Results), different reducing conditions were explored. In some experiments dithiothreitol (1 mM) was used in place of glutathione and found to be more efficient (see Results). After five hr (and formation of species

B-D; see Results Chapter 4 and Figure 4.1), the cuvette was uncovered in order to allow the

48 entry of oxygen and final measurements were again taken at regular intervals over several hr to reveal the final oxidized adduct.

2.2.7 ATS Activation NMR Studies

Reaction mixtures were typically made to the following final concentrations: FPIX (0.375 mM), glutathione (1.88 mM) and ATS (0.375 mM). To study the effect of partner antimalarial drugs on the reaction between ATS and FPIX, an equimolar amount (0.375 mM) of the partner drug was added to this mixture. The reaction mixtures were kept at 37oC and buffered with phosphate buffer to pH 7.0. Reactions were monitored by 1H NMR spectroscopy using a Varian

Unity INOVA 500 MHz NMR Spectrometer (Agilent). The 3-CH3 resonance of ATS before and after reaction with Fe(II)PIX was monitored in order to measure the rate of reaction between the two reagents, as this methyl is lost upon formation of an ATS-FPIX adduct. NMR resonances were obtained every 30 min for 16 hours. This experiment was performed in the absence and presence of additional quinoline antimalarial drugs and in the presence of increasing equivalents of CQ (0:1, 1:1, 1:2, and 1:4 CQ:FPIX).

Several of these reaction mixtures were also monitored by mass spectrometry using mixtures of hemin chloride, glutathione, ATS, and either QN or epiQN (1:5:1:1 molar ratio) in

H2O at pH 7.0. Solutions were diluted in 1:1 MeOH:H2O and the final mixtures were subjected to electrospray ionization mass spectrometry (ESI-MS), in the positive ion mode, using a Varian

500 ion trap mass spectrometer (Agilent). The electrospray ionization source was operated at 5 kV with N2 as the nebulizing gas. First-order ESI mass spectra were recorded in the mass range m/z 900–1400. The drug-heme solutions were introduced into the mass spectrometer by a syringe pump (Hamilton), employing a 1 mL syringe at a constant flow rate of 10 µL/min.

49 2.2.8 Synthesis of Art Drug Probes

2.2.8.1 Synthesis of Biotinylated Artesunate Probe

ATS (0.80 mmol) was dissolved in 750 µL dichloromethane. Deionized water (10 mL)

was added to the solution, and the mixture was pipetted vigorously, heated to 50 C, and 5 mg of

the phase transfer catalyst tetra-butyl ammonium bromide was added. After 1 hr, N-(3-

Dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC·HCl) (0.71 mmol), 4-

Dimethylaminopyridine (DMAP) (0.71 mmol), and Pentylamine-Biotin (0.80 mmol) were

added. The mixture was stirred overnight at 40°C, then acidified with 15 mL of 6 M HCl and

washed with DCM, saturated NaHCO3 and brine (1x each). The aqueous layer was slowly

evaporated and the pure final product formed as tan crystals. Product was purified to remove

unreacted starting materials via column chromatography over silica gel (30% EtOAC/ Hexanes).

13 C NMR (CDCl3 100 MHz; m = multiplet) δ 12.5, 14.9, 19.5, 22.9, 23.3, 23.5, 24.3, 25.5, 26.4,

26.6, 27.9, 28.0, 29.3, 34.0, 35.6, 37.1, 38.4, 39.5, 39.7, 39.9, 40.1, 40.2, 43.3, 52.3, 55.6, 58.3, 6

0.5, 62.4, 107.2, 157.6, 160.4, 165.5, 179.1.

H O O O H O O O O N OH O O HN NH O H O O O O EDC·HCl H H HN NH DMAP N N S O H H O O N NH2 S O (1)

Scheme 2.1. One-pot synthesis of biotinylated artesunate probe

2.2.8.2 Synthesis of Alkynyl Dihydroartemisinin Probe

DHA (1 g) was dissolved in 25 mL DCM to 0.15 M. The mixture was placed on a dry ice

bath (-50°C), allowed to cool, and then stirred for 30 min. (Trimethylsilyl)-acetylene (1 mL) and

50 BF3 (1.2 mL) were added to the mixture and stirred for 6 hr. As the dry ice melted, additional ice was added to maintain a temperature ~ 0°C. The mixture was washed with water / DCM. The organic layer was collected, dried with MgSO4, moved to a weighed round-bottom flask and concentrated under reduced pressure to yield an oil that was then further purified using column chromatography over silica gel ( 20% EtOAC/ Hexanes) to yield the final product as a light

13 brown oil in 65% yield. C NMR (CDCl3 100 MHz; m = multiplet)

δ 12.3, 16.3, 19.9, 21.9, 23.0, 23.1, 23.5, 23.9, 24.3, 26.2, 31.0, 33.6, 33.9, 34.7, 34.9,

35.1, 39.4, 41.7, 44.5, 44.6, 48.0, 82.3, 91.2, 91.4, 91.9, 92.5, 99.6, 100.0, 103.4, 104.6,

110.5 (m) , 209.8.

H H H O O O Si Ac2O, DMAP O O O O O O H H BF3 OEt2, CH2Cl2 H O O O -78 C to r.t.

OH OAc

(2)

Scheme 2.2. Synthesis of terminal alkynyl artemisinin

51 H O H O O H O N O N Fluorophore H N O H 2 H O 1 O CuL O n-1 O H [L Cu]+ O n H O N N N 5 Fluorophore Ln-2Cu N H N Fluorophore N O O H O H 3 N CuLn-1 O O N N Fluorophore H O 4 N N CuL N n-2 Fluorophore

Scheme 2.3. Addition of fluorophore to clickable alkynyl artemisinin. The terminal alkyne forms an activated copper acetylide (1) which is attacked by an azide nitrogen (2). Formation of a 6-membered metallocycle (3) is followed by contraction of the ring (4) and release of the copper catalyst, yielding the triazole product (5) [Himo et al., 2005].

2.2.9 IC50 Assays to Determine Biological Activity of Probes

Antiplasmodial growth inhibitory (IC50) activity was determined for HB3 and Dd2 strains as previously described. [Bennett et al., 2004] This cytostatic assay involves continuous growth in the presence of low concentrations of drug for 72 hours so that all parasites have undergone a full replication cycle, and the assay can then titrate drug induced changes in the rate of parasite growth. The test compounds, i.e. DHA, ATS, as well as Art derived drug probes were all dissolved in DMSO. For this assay, parasite culture is prepared at 4% hematocrit and 1%

52 parasitemia, i.e. a low enough parasitemia to be able to grow untouched for three days without additional media. 100 µL of culture is then added to 100 µL of serially diluted drug in each well of a sterile 96-well clear-bottom plate. Plates are transferred to an airtight chamber and purged with 5% CO2 / 5% O2 / 90% N2 gas mixture for 3 min and then sealed. After incubating the plate at 37 °C for 72 hours, 50 µL of 10X Sybr Green I dye (Thermo Fisher Scientific (Waltham,

MA)) was diluted into complete media from a 10,000X DMSO stock and added to each well.

The plate was then incubated for an additional hour at 37 °C to allow the Sybr Green dye to react with parasite DNA. Fluorescence for each well the 96-well plate was then measured at 538 nm using a Spectra Gemini plate reader (Molecular Devices; Sunnyvale, CA) Standard curves of known parasitemia vs. fluorescence were prepared immediately prior to addition of Sybr Green and plate analysis. IC50 values were obtained by curve fitting the raw data to % growth vs. log[drug] using SigmaPlot 12.0.

2.2.10 Subcellular Fluorescence Imaging of Art Drug Probes

2.2.10.1 Fluorescence Imaging of Biotinylated Artesunate Probe

iRBCs harboring synchronized trophozoite Dd2 parasites (100 uL) were centrifuged at

1800 rpm for 1 min and resuspended in 100 µL of either 100 nM probe, 100 nM ATS, or PBS

(negative control). Cells were incubated for 30 min at 37°C, then washed 3x with 1X PBS.

Parasites were fixed with 4% paraformaldehyde, 0.004% glutaraldehyde, and incubated for 30 min at 37°C. Samples were washed 3x with 1X PBS. The mixture of paraformaldehyde and glutaraldehyde is an additive fixation solution, which creates covalent chemical bonds between proteins, preserving their natural structure within cells. The final samples were resuspended in

500 µL PBS and a 200 µL aliquot was added to a poly-L-lysine coated #1.5 glass coverslip, which was then incubated at 37°C for 5 min. Non-adherent cells were removed by washing 3x

53 with 1X PBS. Cell membranes were permeabilized with 0.1% Triton-X at 37°C for 10 min, and cells were washed again 3x with 1X PBS. The sample was then diluted with 150 µL AlexaFluor

647-conjugated streptavidin (1:200 dilution) and incubated for 45 min. Coverslips were washed

5x with 1X PBS-Tween 20, mounted onto a glass slide, dried and imaged.

2.2.10.2 Fluorescence Imaging of Alkynyl Dihydroartemisinin Probe

In order to view the alkynyl DHA probe within HB3 parasites via fluorescence microscopy, parasites were photolabeled with the alkynyl DHA probe by UV-irradiation to “click” the alkynyl DHA to an azido fluorophore (Scheme 2.3). Drug (100 nM) was incubated in dark for 15 min and then in UV (254 nm) light for 15 more min using a hand-held lamp (Spectroline model

ENF-280C, 115 V, 60 Hz, 200 mA; 8W bulb emitting approximately 500 µW/cm2 approximately 10 cm from the bulb surface). Cells were fixed with formaldehyde and washed with PBS to remove non-adherent cells as described above (section 2.2.10.1). The samples were then added to coverslips and cell membranes were permeabilized with Triton-X as described above. Probe-treated cells were then coupled to azide-labeled Alexa Fluor 488. Coupling was performed using 200 µM copper(II) sulfate, 200 µM bathophenanthroline disulfonic acid, 1 mM

DTT, and 1:200 dilution of azide-labeled Alexa Fluor 488. Cells were then adhered to poly- lysine-coated #1.5 coverslips for imaging.

DIC data were collected with type A immersion oil (η= 1.515), a halogen lamp for transmittance illumination, and an LP520 filter for transmittance filtering. Spacing in z- was set to 0.2 µm. Transmittance was focused until hemozoin within the parasite DV was clearly visible and presented the largest diameter possible so that the defined z-axis focal plane was within the

DV. This z-axis position was defined as “0 µm”. A series of 31 DIC images or “Z-stacks” were acquired in successive 0.2 µm z-axis displacements within parasite between the -3 µm to +3 µm

54 positions. For acquisition of fluorescent images, the power was set to 100% with 500 ms exposure with excitation/emission wavelengths of 642 nm / 700 nm.

AutoQuant Software was used to deconvolve images. For this process, the microscope records a series of optical sections (stack) along its z-axis and data from each section is recorded by the detector and stored. Afterward, the sections are combined to form a three-dimensional object. The image starts out blurred or “convoluted” as a result of the point spread function, which is based on an infinitely small point source of light originating in the specimen. Only a fraction of the light emitted by this point is collected and a dimensionally accurate image cannot be created. Applying a coordinate system, where the x and y axes are parallel to the focal plane and z is perpendicular, gives the point spread function an hourglass shape. The raw data gives images stretched along the z-axis because the resolution in the z-direction is much smaller than that of the x or y direction. In order to correct for this, an adaptive point spread function is applied which is dependent on the numerical aperture of the microscope, the objective, medium, and wavelength of light used. A point spread function from a previously imaged single fluorescent bead can also be used if measured using the same conditions as the current experiment. Deconvolution optimizes the image through repetition. Each iteration, it updates the object estimate and also the point spread function. Iterative deconvolution (more specifically, bind constrained iterative deconvolution) corrects for the inconsistent optical path: as the z-axis moves up and down, this method takes into account how the PSF changes throughout sample.

55 CHAPTER 3. QUANTIFICATION OF FREE FERRIPROTOPORPHYRIN IX HEME AND HEMOZOIN

FOR ARTEMISININ SENSITIVE VS DELAYED CLEARANCE PHENOTYPE PLASMODIUM

1 FALCIPARUM MALARIAL PARASITES

3.1 Introduction

For more than 100 years, conversion of ferriprotoporphyrin IX (FPIX) heme to what is now known to be crystallized hemozoin (Hz; also known as malaria pigment) during intraerythrocytic development of malarial parasites has been an intense area of study. Hemozoin formation is essential to parasite physiology and for parasite development within the human host and is a key molecular target for antimalarial drug therapy.

Of the >160 species of Plasmodium malarial parasites known to exist, at least five infect humans, with Plasmodium falciparum being the most lethal. Once injected via the bite of an infected female Anopheles mosquito, P. falciparum parasites quickly localize to the liver where they invade hepatocytes, divide asexually, and are released approximately 2 weeks later as merosomes harboring large numbers of viable merozoites [Soulard et al., 2015]. Merozoites disseminate within the blood and quickly invade red blood cells (RBCs) where they then differentiate from ring stage to trophozoite and finally to schizont. Asexual division again occurs at the schizont stage; mature schizonts lyse RBCs, and new merozoites are released into the bloodstream where they then again infect RBCs. Via signals that are poorly understood, some ring stage parasites are committed to differentiation to gametocytes instead of new merozoites.

Gametocytes, when taken up during a mosquito blood meal, differentiate further within the mosquito where they may undergo sexual reproduction.

Among the many fascinating characteristics of intraerythrocytic parasite differentiation,

1Previously published as: Heller, L. E. and Roepe, P. D.* (2018) Quantification of Free Ferriprotoporphyrin IX Heme and Hemozoin for Artemisinin Sensitive vs Delayed Clearance Phenotype Plasmodium falciparum Malarial Parasites. Biochemistry 57, 6927-6934.

56 the parasite must digest most red blood cell hemoglobin (Hb) to both make room for rapid cellular growth and to provide amino acids for heightened metabolism. This digestion occurs within a specialized lysosome-like organelle known as the digestive vacuole (DV) that is most clearly visible at the trophozoite stage but is initially formed at the ring stage of parasite red blood cell development. The toxic byproduct of parasite Hemoglobin digestion, free FPIX heme, must be converted to nontoxic crystalline hemozoin for the parasite to grow [Francis et al.,

1997]. Several classes of antimalarial drugs, including the well- studied quinoline methanols, amino quinolines, and artemisinin (Art)-based drugs, inhibit FPIX to hemozoin conversion by multiple mechanisms that include binding to one or more forms of free FPIX, adhesion to growing faces of the hemozoin crystal, and perhaps others (L.E.H. and P.D.R., unpublished)

[Gorka et al., 2013; Kuter et al., 2016; de Villiers et al., 2012; Olafson et al., 2017].

Quantifying parasite Hemoglobin digestion can be done by quantification of parasite hemozoin produced from Hemoglobin. From such measurements, calculations of precisely how much Hemoglobin is digested by a parasite vary from ~50 to ~90% of available Hemoglobin

[Asawamabasakda et al., 1994; Ginsburg et al., 1998; Zhang et al., 1999; Gligorijevic et al.,

2006a]. The chemistry of formation of hemozoin from FPIX that is released during Hemoglobin catabolism is dependent on a number of variables. Although many studies have examined hemozoin formation in vitro, few studies have quantified hemozoin production within live parasites at different stages of parasite development, and none to the best of our knowledge have quantified the ratio of free FPIX to crystallized hemozoin at multiple parasite stages. However, such information is important for understanding the pharmacology of antimalarial drugs and also for understanding antimalarial drug resistance phenomena. Several studies have shown that altered FPIX−hemozoin conversion and altered drug− FPIX binding are likely related to

57 resistance to quinoline-based antimalarials [Gligorijevic et al., 2006a; Gligorijevic et al., 2008;

Leed et al., 2002]. Perhaps not coincidentally, several other studies have found altered DV physiology and DV morphology associated with parasite chloroquine resistance (CQR)

[Dzekunov et al., 2000; Bennett et al., 2004; Gligorijevic et al., 2006b; Abu Bakar et al., 2010;

Pulcini et al., 2015]. These alterations in DV physiology likely influence FPIX−hemozoin conversion as well as drug−FPIX interactions, thereby contributing to CQR and possibly other antimalarial drug resistance phenomena [Gorka et al., 2013].

Interestingly then, a recent report found evidence of altered kinetics of intraerythrocytic development for P. falciparum isolated from patients exhibiting a “delayed clearance phenotype”

(DCP) upon administration of Art combination therapy (ACT). Specifically, for DCP parasites relative to the control, the intraerythrocytic ring stage was found to be extended and the trophozoite stage shortened [Hott et al., 2015]. DCP parasites are often termed “Art resistant”

(ArtR) malarial parasites even though they do not show easily measurable shifts in Art IC50 as observed for all other well-known examples of antimalarial drug resistance (see Chapter 4 and

Heller et al., 2018b). As explored in more detail in the accompanying paper (see Chapter 4 and

Heller et al., 2018b), a DCP indicates the altered potency of Art-based drugs at primarily the ring stage of parasite red blood cell development, but without an easily detectable shift in drug IC50.

Regardless, because FPIX may activate prodrug forms of Art and its relatives and because altered cell cycle kinetics might predict an altered abundance of FPIX that would impact Art activation at specific times during parasite development, we endeavored to precisely quantify

FPIX and hemozoin versus the time of intraerythrocytic development for control versus DCP malarial parasites.

One well-defined laboratory model for DCP parasites consists of Pfk13 gene-edited

58 strains [Straimer et al., 2015]. Using these strains, optimized heme and hemozoin extraction protocols, and liquid chromatography−mass spectrometry (LC−MS) methods, we quantify FPIX and hemozoin abundance for ring, trophozoite, and schizont stages of control versus DCP P. falciparum. As I describe further in the accompanying manuscript (see Chapter 4 and Heller et al., 2018b), these data have important implications for understanding Art drug molecular pharmacology and the molecular mechanism of evolving Art resistance.

3.2 Results

Recently, Kyle and colleagues analyzed the growth of two P. falciparum laboratory strains established from parasite isolates taken from patients that exhibited a DCP [Hott et al.,

2015]. In this study, growth of DCP strains was compared to that of a non-isogenic laboratory control strain isolated much earlier (strain W2) [Oduola et al., 1988] that is highly resistant to chloroquine and other drugs but is not a DCP parasite. Recently, isogenic strains that harbor either the wild type or DCP-associated mutant Pfk13 have become available [Straimer et al.,

2015]. In these models, one mutant PfK13- expressing strain was constructed by reverse genetic experiments using a host strain isolated from a non-DCP patient, and another strain expressing wild-type PfK13 was constructed by reverse genetics using a strain cultured from a DCP isolate

[Straimer et al., 2015]. Thus, these strains permit isolation of DCP features that are solely due to mutant PfK13 protein, which has been shown to cause DCP [Straimer et al., 2015; Ariey et al.,

2014; Cheeseman et al., 2012; Takala-Harrison et al., 2013]. We analyzed the kinetics of intraerythrocytic development for these strains (Figures 3.1-3.3).

Figure 3.1. Giemsa smears of live parasites across the life cycle. Schematics highlight the successive stages of the CamWT life cycle at increasing hours post synchronization.

Figure 3.2 Kinetics of intraerythrocytic development for isogenic control strain versus delayed cleanrance phenotype parasites. Left: isogenic control strain CamWT; Right: mutant strain CamWT-C580Y. Rings: black, trophozoites: red, schizonts: green. Reproduced from [Heller et al., 2018a] without permission.

Figure 3.3 Kinetics of intraerythrocytic development for additional control and delayed cleanrance phenotype strains. Left: isogenic control strain Cam3.11-REV; Right: mutant strain Cam3.11-R539T. Rings: black, trophozoites: red, schizonts: green. Reproduced from [Heller et al., 2018a] without permission.

Similar to previous work, [Hott et al., 2015] we find that the ring stage of intraerythrocytic development is extended and that the trophozoite stage is shortened for DCP parasites (e.g., strain Cam580Y, Figure 3.2) relative to their isogenic non-DCP control [strain

CamWT, Figure 3.2; note the lengthening of the ring stage (black circles) versus the shortening of the trophozoite stage (red circles) (Figure 3.2)]. Similar results were found when comparing

61 DCP strain Cam3.11R539T to its isogenic non-DCP control strain (Cam3.11R539Trev; data not shown). Remarkably, however, the sum of ring extension times for the DCP parasites

(approximately +7 h relative to the control) and the trophozoite shortening time (approximately

−6 h relative to the control) is close to zero such that the overall intraerythrocytic cycle time

(approximately 48 h) remains constant relative to that of the isogenic non-DCP control.

Again, the trophozoite stage of parasite development is thought to be the stage at which the parasite digests most red blood cell Hemoglobin. The fact that wild-type intraerythrocytic P. falciparum digests most of the 5 mM Hemoglobin present in the red blood cell primarily during the trophozoite stage and thereby expands the parasite cellular volume >25-fold in ~18 h (see

CamWT, Figure 3.2, and Gligorijevic et al., 2006a) is remarkable enough in and of itself, but we find the possibility that a DCP parasite might accomplish the same feat in approximately two- thirds of the time (Figures 3.2 and 3.3) to be particularly surprising. Notably, however, the sum of ring and trophozoite times for DCP versus non-DCP controls is the same (approximately 35 h), suggesting that if ring stages of the parasite also catabolize Hemoglobin and if DCP parasites are somehow able to distribute that catabolism across both ring and trophozoite stages more evenly than is the case for non-DCP parasites, then overall parasite red blood cell development to schizogony might not be compromised for DCP parasites. In such a scenario, however, the kinetics of Hemoglobin catabolism (and release of free FPIX as a consequence of that catabolism) would obviously be perturbed. Although a few reports showing ring stage digestion of Hemoglobin have been published [Abu Bakar et al., 2010; Elliott et al., 2008], it has been widely assumed that obligate parasite Hemoglobin digestion occurs primarily during the trophozoite stage of parasite development. To the best of our knowledge, no direct measurements

62 that quantify FPIX and hemozoin in parallel at the ring stage relative to the trophozoite stage have previously been performed.

That is, a convenient method for monitoring both the extent of Hemoglobin catabolism and the release of free FPIX within the P. falciparum DV is to monitor the formation of crystalline Hemozoin from free FPIX heme. Our earlier attempts at kinetic resolution of hemozoin formation included spinning disk confocal quantification of hemozoin within single live parasites [Gligorijevic et al., 2006a]. Although useful, this method does not allow simultaneous quantification of free FPIX heme. If some degree of Hemoglobin catabolism does indeed occur during the ring stage of development as suggested [Abu Bakar et al., 2010; Elliott et al., 2008], then the amounts of free FPIX heme released from Hemoglobin during this stage would be expected to be small. We therefore optimized LC−MS methods to detect low concentrations of FPIX. Figure 3.4A shows a representative spectrum obtained for 100 nM FPIX dissolved in LC−MS grade MeOH with 0.1% formic acid.

63 616.2 35 557.2 30

20 498.2

10 Intensity (kCounts) Intensity

0 400 500 600 700 800 900 m/z

Figure 3.4A Mass spectrum of pure ferriprotoporphyrin IX heme. Solution consisted of 100 nM Hemin-Cl dissolved in methanol with 0.1% formic acid, diluted from a stock solution of 2 mM Hemin-Cl prepared in 0.1 M NaOH. Reproduced from [Heller et al., 2018a] without permission.

N N N -C2H3O2 N -C2H3O2 N N Fe Fe N N Fe N N N N

O OH OH O O OH m/z = 616 m/z = 557 m/z = 498

Figure 3.4B Mass spectrometry fragmentation pattern of pure FPIX heme. Basic fingerprint seen previously in Pashynska et al., 2004. Reproduced from [Heller et al., 2018a] without permission.

As published elsewhere [Pashynska et al., 2004], clear peaks at m/z 616.2, 557.2, and

498.2 characteristic of FPIX and common breakdown products (Figure 3.4B) are easily

64 observed. As shown in Figure 3.5, careful analysis of FPIX solutions at various concentrations permits reliable quantification of FPIX to nanomolar levels using comparison to a linear standard curve (Figure 3.5).

Figure 3.5 Mass spectrometry calibration curve for free heme using pure hemin-Cl. R2 = 0.983. Reproduced from [Heller et al., 2018a] without permission.

Similarly, FPIX that has been incorporated into crystallized hemozoin can be quantified via these MS methods upon NaOH solubilization of crystalline hemozoin to liberate FPIX [Fitch et al., 1987]. Hemozoin is a crystal of FPIX reciprocal or “head-to-tail” dimers [Pagola et al.,

2000] that have predicted m/z values near 1230. Figure 3.6 shows MS data for FPIX after solubilizing hemozoin that was synthesized in vitro according to procedures published elsewhere

[Gorka et al., 2013].

Figure 3.6 Mass spectrum of pure heme solubilized from β-hematin (synthesized in vitro) using NaOH. Both FPIX monomer (616.4) and reciprocal dimer peaks at 1253.3 and 1231.3 [2*FPIX – 2H, + Na] and [2*FPIX - H], respectively are observed [Pashynska et al., 2004]. Reproduced from [Heller et al., 2018a] without permission.

The prominent FPIX peak at m/z 616 is again seen (Figure 3.4A) because NaOH solubilization of hemozoin under these conditions hydrolyzes most FPIX dimers to the FPIX monomer [m/z 616.2 (Figure 3.6)]. However, residual FPIX reciprocal dimer peaks are also observed at m/z 1231.2 and 1254.2. In addition to these LC−MS methods, if sufficient hemozoin is present, FPIX solubilized from hemozoin can be easily quantified via ultraviolet−visible

spectroscopy as previously described (See Methods, Figure 2.4) [Slater et al., 1992].

Using these principles and synchronized bulk parasite culture, we quantified both free

FPIX heme and hemozoin (as hydroxide-solubilized FPIX) from parasites versus the time of red blood cell development. Figure 3.7A shows representative LC−MS data for free FPIX isolated from iRBCs infected with CamWT strain parasites 32 h post-invasion, and Figure 3.7B shows

LC− MS data for NaOH-solubilized hemozoin purified from CamWT strain parasites 37 h post-

66 invasion (see Methods). With the exception of a minor peak corresponding to small amounts of lipid that are also extracted from the parasite with these methods [m/z 760.6 (Figure 3.5A)],

LC−MS data for free FPIX and hemozoin FPIX extracted from parasites are quite similar to data for pure compounds prepared in vitro (compare Figure 3.4A to Figure 3.7A and Figure 3.6 to

Figure 3.7B, respectively). Figure 3.7C shows LC−MS data for a control CHCl3 extract from non- infected RBCs prepared in the same fashion as parasites released from infected RBCs (see

Methods) and reveals that no free FPIX heme is extracted from RBCs under these conditions.

300 616.3

250

200

150 557.2

100 Intensity (kCounts) Intensity

50 498.3 760.6

0 400 450 500 550 600 650 700 750 800 850 900 m/z

Figure 3.7A. Mass spectrum of free heme isolated from synchronized CamWT trophozoites. Note signature FPIX peak at m/z 616.3. Common mass spectrometry breakdown products are also again visible (compare to Figure 3.2A). Higher m/z = 760.6 represents minor co - extracted phospholipid [Pisciotta et al., 2007]. Reproduced from [Heller et al., 2018a] without permission.

67 200 616.1

180

160

140

120

100 1253.3

Intensity (kCounts) Intensity 60

40 1231.3

659.151 20

0 400 500 600 700 800 900 1000 1100 1200 1300 m/z

Figure 3.7B Mass spectrum of free heme solubilized from isolated parasite hemozoin. Note the m/z 616.1 peak (FPIX monomer) as well as FPIX reciprocal dimer peaks at 1253.3 and 1231.3 [2*FPIX – 2H, + Na] and [2*FPIX -H], respectively [Pashynska et al., 2004]. Compare to Fig. 3.4. Reproduced from [Heller et al., 2018a] without permission. 150

120

60 Intensity (kCounts) Intensity

0 400 450 500 550 600 650 700 750 800 850 900 m/z

Figure 3.7C Mass spectrum of chloroform extracts of non infected red blood cells. Extracts prepared in the same fashion as those for infected red blood cells. No soluble heme is visible. Reproduced from [Heller et al., 2018a] without permission.

68 Because the intraerythrocytic development of DCP parasites is kinetically distinct from that of controls [Hott et al., 2015] (cf. Figures 3.2 and 3.3), we next quantified the abundance of free FPIX and hemozoin FPIX versus the time of intraerythrocytic development for control versus DCP parasites (Figures 3.8, 3.9 and 3.10). As shown, non-DCP CamWT parasites (Figure

3.8, black circles) show peak abundance of free FPIX coincident with the trophozoite stage of development (28−34 h), with very low levels at other stages, consistent with maximum production of hemozoin from free FPIX at the trophozoite stage observed previously [Zhang et al., 1999; Gligorijevic et al., 2006a; Xie et al., 2016]. In contrast, isogenic DCP Cam580Y parasites exhibit a much broader free FPIX peak centered at the trophozoite stage, with higher levels of FPIX apparent at the ring and schizont stages relative to CamWT [Figure 3.8, black filled circles (control strain CamWT) to open circles (DCP strain Cam580Y)].

Figure 3.8 Free heme abundance across the parasite intraerythrocytic life cycle. Black circles: CamWT, open circles: CamWT-C580Y. Data represent at least two independent experiments (two independent parasite culture growths) with two replicates (two independent FPIX measurements) of each for 4 determinations in total. Arrows represent time points where data for strains Cam3.11-R539T (delayed clearance phenotype, gray circles) and Cam3.11- R539T-Rev (isogenic control, green circles) were also collected. All stage - dependent differences between either delayed clearance phenotype parasite and the relevant matched isogenic control at points with green arrows (e.g., "CamWT-R" vs "C580Y-R", "Rev-S" vs "R539T-S", etc.) are statistically significant (p < 0.05). Reproduced from [Heller et al., 2018a] without permission.

Hemozoin was also quantified versus time for these strains (Figures 3.9 and 3.10). As shown, in spite of the very different abundance of free FPIX versus time during intraerythrocytic development, the total amount of hemozoin produced by the two strains is more similar. The kinetics of hemozoin formation differs for the two strains, with DCP strain Cam580Y (Figure

3.10, open circles) showing faster hemozoin production relative to matched isogenic non-DCP strain CamWT (Figure 3.10, filled circles). Maximum rates of hemozoin production for both strains are found at times that correspond approximately to the maximum abundance of free

FPIX [28− 32 h (compare Figure 3.8 vs Figure 3.10, see also Figure 3.11)].

Figure 3.9. Quantifying hemozoin by UV-Visible absorbance spectroscopy. Hemozoin was isolated after washing parasite pellets with 0.1 M NaHCO3 and dissolving remaining pellets in 0.1 M NaOH. Time points are as follows, where HPI represents number of hours post invasion: Blue: 3 HPI; Cyan: 9 HPI; Red: 22 HPI; Orange: 28 HPI; Green: 30 HPI; Black: 46 HPI.

400

300

200

100 [FPIX from Hz] (mM) Hz] from [FPIX

0 10 20 30 40 50

Time (Hours Post Synchronization)

Figure 3.10 Levels of hemozoin across the parasite intraerythrocytic life cycle. Hemozoin purified from strains CamWT (closed circles) and Cam580Y (open circles). Data represent at least two independent experiments (two independent parasite culture growths) with two replicates (two independent FPIX measurements) of each (4 determinations in total). Reproduced from [Heller et al., 2018a] without permission.

Finally, to test whether differences in free FPIX abundance versus time of intraerythrocytic development are a general characteristic of DCP malarial parasites, we extracted FPIX and hemozoin for an alternate genetically matched set of control (strain

Cam3.11R539Trev) versus DCP (strain Cam3.11R539T) parasites [Straimer et al., 2015]. As shown in Figure 3.8, similar to the comparison between strains CamWT (control, black circles) and Cam580Y (DCP, open circles), DCP strain Cam3.11R539T (light gray circles) harbors significantly reduced levels of free FPIX versus the isogenic control (strain R539TRev, green circles) at the trophozoite stage of development (marked by the upward arrow) but significantly increased levels of free FPIX at the ring and schizont stages of development (marked by the first and second downward arrows, respectively).

3.3 Discussion

The results of this study may be summarized as follows.

(1) The kinetics of intraerythrocytic development for DCP (artemisinin “resistant”) P. falciparum malarial parasites differ from those of matched isogenic control strains. We find that DCP parasites show extended ring stages of development and shortened trophozoite stages of development, with no net increase in the overall intraerythrocytic development cell cycle time, similar to earlier observations from the Kyle laboratory [Hott et al., 2015].

(2) For the first time, free parasite FPIX and FPIX liberated from parasite hemozoin, both isolated in parallel from synchronized live P. falciparum culture, are quantified for all parasite stages using perfected LC−MS methods.

(3) Along with altered intraerythrocytic development, for the first time we find that DCP parasites show altered levels of free FPIX relative to the matched isogenic control throughout the

72 intraerythrocytic cell cycle as well as slightly faster and elevated levels of hemozoin production from free FPIX. Interestingly, the level of free FPIX is lower for DCP parasites at the trophozoite stage but higher at the ring and schizont stages of development.

These data have important implications for understanding both parasite intraerythrocytic development and the mechanism(s) behind the DCP, which is believed to be a harbinger of evolving Artemisinin resistance.

It has been known for some time that easily measurable free FPIX and copious hemozoin are present at the trophozoite stage of parasite development. Our data verify this to be the case but also show that measurable levels of free FPIX heme are present for P. falciparum at the ring and schizont stages. Correspondingly, low, but measurable, levels of hemozoin FPIX are found at the late ring−early trophozoite boundary. With these methods, which rely on extraction of bulk parasite culture after visual definition of (mixed) parasite stages versus time, we cannot unequivocally determine whether hemozoin is produced for late rings, early trophozoites, or both, but on the basis of previous work by others and what is currently known about the hemozoin crystallization process [Gorka et al., 2013; Gligorijevic et al., 2006a; Abu Bakar et al.,

2010; Elliott et al., 2008], it seems likely that small amounts of hemozoin crystallization are initiated during the late ring stage.

Interestingly, levels of free FPIX differ for two different strains of DCP parasites relative to their isogenic controls, across all stages of parasite development. DCP free FPIX levels are higher (p < 0.05; Student’s t test) at ring and schizont stages relative to that of the control and conspicuously lower at the trophozoite stage. As explored in the following chapter (see chapter 5 and also Heller et al., 2018b), these data have important implications for the availability of activated Art drugs at various stages of parasite development.

73 It is also interesting to note that the time at which the highest abundance of free FPIX heme is found for strain CamWT (31 h post-infection) overlaps precisely with the midpoint for hemozoin production (Figure 3.11).

Figure 3.11. Overlays of free heme and hemozoin production. Left: CamWT, Right: CamWT-C580Y. Green: % free heme, normalized to max [heme]; Blue: % hemozoin, normalized to max [hemozoin] levels.

In contrast, for DCP parasites (e.g., strain Cam580Y), the highest abundance of free FPIX heme occurs at the same absolute time point (31 h), but this time does not overlap with the midpoint of hemozoin production, which occurs conspicuously earlier, suggesting that the dynamics of FPIX to hemozoin conversion differ for DCP parasites.

Altered FPIX and hemozoin abundances are found for two different DCP strains that were created by reverse genetic experiments that replaced the wild-type Pfk13 gene with a mutant gene that expresses a PfK13 580Y mutant (CamWT vs Cam580Y) or were cultured from a DCP isolate, with the isogenic control then created by replacing mutant Pfk13 found in the isolate with wild-type Pfk13 (Cam3.11R539T vs Cam3.11R539- Trev) [Straimer et al., 2015].

Notably, mutation of PfK13 to either 580Y PfK13 or 539T PfK13 causes similar changes in free

74 FPIX heme levels. Examining strains harboring other PfK13 changes, particularly any that are found to confer altered degrees of DCP, should prove to be interesting.

These experiments link mutant PfK13 function to changes in free FPIX heme abundance during the parasite intraerythrocytic cell cycle. These changes in FPIX are likely a consequence of altered cell cycle kinetics, meaning the extended ring stage and shortened trophozoite stage of development, as previously noted by Kyle and others [Hott et al., 2015; Mok et al., 2015].

These changes in cell cycle kinetics might be mediated by an as yet to be identified cell signaling function for PfK13. PfK13 shows some homology to human KLHL proteins involved in ubiquitin-based protein degradation [Pei et al., 2018; D’Angioelella et al., 2013]. Because the levels of some cyclin proteins that regulate cell cycle kinetics are regulated by ubiquitin- mediated pathways [Pei et al., 2018; D’Angioelella et al., 2013], perhaps mutations in PfK13 alter parasite intraerythrocytic cell cycle kinetics through effects on one or more parasite cyclins.

Assuming some catabolism of Hemoglobin at the ring stage [Abu Bakar et al., 2010; Elliott et al., 2008], a longer ring stage instigated by dysregulation of the intraerythrocytic cell cycle might predict that higher levels of free FPIX heme might accumulate for rings as is indeed observed.

This scenario assumes that the rate of hemozoin production remains similar for DCP rings. A shorter trophozoite stage might predict lower levels of free FPIX heme relative to control as is also observed. Another hypothesis for DCP-associated mutant PfK13 function that has been offered is that the protein affects phosphatidyl 3′-kinase activity [Mbengue et al., 2015]. How altered PI3P levels might then affect vesicle-mediated Hemoglobin catabolism and free FPIX levels is yet to be defined, but one recent study suggests that parasite PI3P levels influence vesicle traffic [Bhattacharjee et al., 2018] These hypotheses and others that might explain how

PfK13 mutations affect free FPIX heme levels merit further study.

75 CHAPTER 4. DIHYDROARTEMISININ-FERRIPROTOPORPHYRIN IX ADDUCT ABUNDANCE IN

PLASMODIUM FALCIPARUM MALARIAL PARASITES AND THE RELATIONSHIP TO EMERGING

1 ARTEMISININ RESISTANCE

4.1 Introduction

Plasmodium falciparum malaria remains a threat to more than half of the world’s population and kills more than 0.5 million annually. Antimalarial drugs have been, and remain, the most effective treatment for malaria. Because resistance to quinoline- based and antifolate antimalarial drugs compromises their use in many areas of the globe, the currently recommended treatments for P. falciparum malaria are several available “artemisinin combination therapies”

(ACTs). These combine the use of an artemisinin (Art)-based drug (e.g., the “parent” drug) with one of several possible “partner” drugs. The most commonly used ACT is Coartem, which is comprised of artemether (ATM, the ether derivative of the natural product Art) and lumefantrine

(LF). Other examples include dihydroartemisinin with piperaquine (DHA/PPQ) and artesunate with mefloquine (ATS/MQ).

Although some data for Art are conflicting, [Lee et al., 1989] all Art-based drugs except one (artemisone) presumably convert to DHA. Rates of conversion vary (e.g., Art vs ATM,

ATS), but conversion is not necessarily metabolic; rather, at least for ATS and ATM, it is known to be spontaneous in plasma and other aqueous media. More data (particularly for Art, which also decays to deoxy-Art under acidic conditions) are needed to better understand relative spontaneous conversion rates. [Lee et al., 1989] In the clinic, a detailed study of the Art:DHA,

ATM:DHA, or ATS:DHA plasma ratios at different times post-ACT treatment has not been

1Previously published as: Heller, L. E., Goggins, E., and Roepe, P. D.* (2018) Dihydroartemisinin-Ferriprotoporphyrin IX Adduct Abundance in Plasmodium falciparum Malarial Parasites and the Relationship to Emerging Artemisinin Resistance. Biochemistry 57, 6935-6945.

76 performed, but available data suggest that, once administered to malaria patients, common ACTs reduce to predominantly DHA and partner drug within approximately 20 min. [Hien et al., 2004]

Currently, ACTs are the only universally effective treatments versus all malaria worldwide. Troublingly, however, a harbinger of Art drug resistance has emerged in southeast

Asia, [Noedl et al., 2008; Dondorp et al., 2009] termed the “delayed clearance phenotype”

(DCP). Shifts in either Art cytostatic potency (quantified as IC50) or cytocidal (parasiticidal) potency (quantified as LD50) [Roepe et al., 2014] have not generally been observed for parasite isolates cloned from malarial patients exhibiting a DCP, although one intriguing report does find

LD50 shifts for ring stages of laboratory- cultured strains of DCP P. falciparum. [Klonis et al.,

2013] DCP is more routinely defined by clinical criteria. [Dondorp et al., 2009; Flegg et al.,

2011; Amaratunga et al., 2012] In brief, patients infected with DCP malaria are typically still cured by ACTs, but the profound ~3 logarithmic drop in parasitemia that typically occurs in ~3 h upon ACT administration requires longer treatment (e.g., 5−6 h to occur). In the laboratory, DCP is quantified in one of several ways, most popularly via a ring stage assay (RSA) that uses very young ring stage parasites (0− 3 h post-invasion) and involves bolus dosing of cultured parasites with higher levels of the Art-based drug (corresponding to those found in human plasma, not to laboratory IC50 levels; ≥700 nM is typically used), typically in the absence of the relevant ACT partner drug. [Witkowski et al., 2013] These assays clearly show that most Art-based drugs have decreased potency versus DCP parasites, [Siriwardana et al., 2016] but whether DCP parasites with reduced ring stage susceptibility to DHA are also resistant to common ACT partner drugs

(e.g., LF, PPQ, and MQR) is often not known. Thus, although often also termed “Art resistance”

(ArtR), DCP actually refers to a spectrum of complex resistance phenomena that continue to evolve.

77 Although the molecular mechanism of DCP is currently unknown, one class of genetic markers for DCP has recently been identified. These are single-nucleotide polymorphisms

(SNPs) in the Pfk13 gene of P. falciparum, leading to single- amino acid substitutions in the encoded PfK13 protein. The function of wild type PfK13 protein is unknown, although one hypothesis is that the protein may play a role in protein folding and/or ubiquitinylation. [Ariey et al., 2014] How the function of mutated PfK13 promotes DCP is unknown. Mbengue et al. recently suggested that PfVps34 (the sole P. falciparum class III phosphatidylinositol 3′-kinase

[Hassett et al., 2017a]) forms a complex with mutant PfK13 and that mutant PfK13 may indirectly affect phosphatidyl 3′-phosphate (PI3′P) production by PfVps34. Via this model, the activity of Art drugs was proposed to be due to noncovalent binding to PfK13, stabilized via hydrogen bond interactions involving an intact Art endoperoxide bridge.

However, to the best of our knowledge, all other studies indicate that the activity of Art- based drugs is dependent upon cleavage of the drugs’ endoperoxide bridge to generate highly reactive carbon-centered radicals that are then capable of alkylating numerous targets within the parasite. [Posner et al., 1992; Asawamahasakda et al., 1994; Robert et al., 2005] Indeed, two recent reports identify many dozens of protein targets for endoperoxide-cleaved, or “activated”,

Art-based drugs, [Ismail et al., 2016a; Wang et al., 2015] but there is only partial overlap between the two sets of proteins identified in these studies (123 protein targets are found in

Wang et al. and 49 in Ismail et al., with 26 overlapping between the two sets). This suggests that alkylation of protein targets within the parasite may be somewhat random. Consistent with these observations, inhibition of PfVps34 by Art-based drugs was recently shown in fact also to require activation of the drug endoperoxide and covalent binding of the drug to PfVps34, with sites of covalent DHA−PfVps34 interaction appearing to be random. [Hassett et al., 2017b]

78 In addition to the PfVps34 target hypothesis, because of the requirement for Fe2+ in catalyzing endoperoxide activation of DHA and other artemisinin-based drugs, it has also been hypothesized that FPIX heme released during parasite Hemoglobin catabolism could be an important target of Art-based drugs. [Robert et al., 2005; Wang et al., 2016; Zhang et al., 2008;

Creek et al., 2008; Zhang et al., 2009; Mercer et al., 2011] That is, the most biologically relevant means of activating DHA and other Art-based drug endoperoxide groups to create carbon-centered radicals capable of alkylating Art targets is via Fe2+-catalyzed Fenton chemistry.

[Posner et al., 1992; Robert et al., 2001] In the malarial parasite, the by far most abundant potential source of Fe2+ is ferriprotoporphyrin IX (FPIX) heme, released during obligate hemoglobin (Hb) catabolism by the intraerythrocytic parasite. FPIX Fe2+ is as potent as free Fe2+ in activating the Art drug endoperoxide bridge. [Hassett et al., 2017b] To the best of our knowledge, free Fe2+ does not exist in eukaryotic cells; however, levels of “labile” iron are 2−5

µM. [Loyevsky et al., 1999; Kakhlon et al., 2002] In contrast, concentrations of iron in the form of FPIX can exceed 4 mM within the malarial parasite DV. [Heller et al., 2018a] Taken together, these observations predict that FPIX is the primary activator of Art - based drugs within P. falciparum.

Also, considered along with the known short lifetime and highly reactive nature of Art - based drug carbon-centered radicals, this also predicts the formation of drug−FPIX covalent adducts during Art - based drug activation by FPIX liberated within the DV of P. falciparum.

Such FPIX-mediated DHA activation and subsequent FPIX−DHA adduct formation can be easily observed in vitro using pure drug, pure heme, and adequate reducing agent to form ferrous

FPIX, but such adducts have not previously been isolated from live P. falciparum parasites. The

Art−heme adduct has however been observed in the blood of Plasmodium vinckei vinckei-

79 infected mice after Art treatment, [Robert et al., 2005] suggesting that adduct could possibly be formed within some rodent malarial parasites.

Art drug alkylation of FPIX would inhibit detoxification of free FPIX to inert hemozoin and would lead to the buildup of toxic drug−FPIX adducts that would then have a variety of additional effects on the parasite. In this paper, we test whether FPIX−DHA adducts are formed within P. falciparum parasites, test whether their abundance varies versus the stage of intraerythrocytic parasite development, and test whether differences in their abundance exist for control versus DCP parasites. The data suggest improved models for Art drug activity versus malarial parasites and for emerging artemisinin resistance.

4.2 Results

Several studies have demonstrated formation of covalent Art drug−FPIX heme adducts in vitro and have provided chemically logical schemes for the overall reaction leading to adduct formation. [O’Neill et al., 2004; Robert et al., 2002a; Robert et al., 2002b] In brief, activation of the Art endoperoxide bridge by reduced FPIX Fe2+ leads to generation of an oxy-centered Art radical that then rearranges to yield an Art carbon-centered radical (most commonly the “C4”- centered radical) that then attacks one of three preferred meso carbons on the tetrapyrrole ring of

FPIX [Robert et al., 2002a] to form a covalent adduct. Spectroscopic evidence for predicted reaction intermediates is rare however. To validate and further illuminate the proposed reaction scheme (Figure 4.1) [O’Neill et al., 2004; Robert et al., 2002a], we monitored reaction of DHA with FPIX heme by UV−vis spectroscopy under conditions that both stabilize FPIX heme in monomeric form and would slow conversion between putative intermediates (so that UV−vis spectra could be better resolved).

A B C

N N Fe N N N N N N Fe3+ Fe2+ O OH N N GSH N N DHA O OH GSSG 2 1 H2C O O O OH O OH OH O OH O Oxy H3C O Fe(III)PPIX Fe(II)PPIX Radical O OH Complex 3

D E

N N N N Final Initial 2+ O2 Fe3+ Fe Oxidized Covalent N N HO N N HO Complex Complex HO 4 HO O O OH OH O O OH O OH OH O OH

Figure 4.1 Intermediates in the production of ferriprotoporphyrin IX - dihydroartemisinin covalent adduct. The first step in the formation of heme – artemisinin adduct is the reduction of Fe(III) heme (A) to Fe(II) heme (B) by glutathione, followed by the addition of dihydroartemisinin (DHA) (2) which instantly forms an iron-oxo intermediate (C). Intramolecular rearrangement (3) gives the covalent complex (D) and final oxidation (4) yields (E). Reproduced from [Heller et al., 2018b] without permission.

Reduction of ferric FPIX with a 5-fold molar excess of glutathione is easily monitored by

UV−Visible spectroscopy (Figure 4.2A,B) with a dramatic sharpening and red-shift of the heme

Soret band (present at 380 nm when FPIX is dissolved in an aqueous DMSO solution) to 419 nm. Upon addition of equimolar DHA, the heme absorbance immediately broadens and blue- shifts to 387 nm as the ferric FPIX−DHA oxo radical dative complex (Figure 4.2C) is formed.

This intermediate is relatively short-lived and rearranges to form the ferrous FPIX−C4−DHA covalent complex (Figure 4.2D) that absorbs maximally near 421 nm (Figure 4.3). NMR data

81 clearly show that drug deacetylation occurs during rearrangement of the oxo radical to the C4 radical and formation of the FPIX−C4−DHA complex (Figure 4.2C vs 4.2D; see also Robert et al., 2005; Robert et al., 2002a; Laurent et al., 2005). Finally, slow re- oxidation of FPIX Fe2+ in complex D results in the final oxidized FPIX−DHA adduct (Figure 4.2E) with a reduced absorbance near 408 nm. Importantly, the first and third steps of the reaction (Figure 4.1) that generate B and D from A and C, respectively, are both highly dependent on pH with rates slower at more alkaline pH (data not shown) due to H+ dissociation associated with GSH/GSSG cycling and DHA carbon−FPIX meso carbon single bond formation, respectively. A more detailed analysis of these pH dependencies will be presented elsewhere.

82 380

A BB N N 3+ Fe FPIX (Fe3+) N N

250 450 650 O OH O OH

419

B N N Fe2+ N N FPIX (Fe2+)

O 250 450 650 OH O OH

387 N N Fe C N N 3+ O OH FPIX (Fe )-oxo-DHA O OH

H2C 250 450 650 O O H3C O O 421 OH D N N Fe2+ N N HO 2+ FPIX (Fe )-C4-DHA HO O 250 450 650 OH O OH O OH

408 E N N Fe3+ N N HO 3+ FPIX (Fe )-C4-DHA HO O 250 450 650 OH O OH OH Wavelength (nm) O

Figure 4.2. Absorbance spectra of ferriprotoporphyrin IX heme after addition of glutathione and dihydroartemisinin. The soret band shifts between the absence (A) versus presence (B) of glutathione, and after the addition of dihydroartemisinin (C-E). Structures corresponding to the observed spectra are shown to the right. Reproduced from [Heller et al., 2018b] without permission.

Figure 4.3. Time dependent reaction between ferriprotoporphyrin IX heme, glutathione, and dihydroartemisinin as monitored by UV-Visible absorbance spectroscopy. Top panel: reduction of Fe(III)PPIX to Fe(II)PPIX by addition of glutathione; Middle: subsequent oxidation immediately upon addition of DHA as an F+3-oxo intermediate is formed; Bottom: eventual reduction as drug complex rearranges to form a covalent adduct.

84 Production of the ultimate FPIX−DHA adduct “E” is easily monitored by LC−MS as well. After reaction of fully reduced ferrous FPIX with equimolar DHA to completion and extraction of the solution using the same protocol developed for extraction of heme from parasites (See Chapter 3) [Heller et al., 2018a], a peak characteristic of the adduct at m/z 881

(Figure 4.4A) is clearly visible.

250 880.9

200

150 881.9

100 Intensity (kCounts) Intensity

0 850 860 870 880 890 900 m/z

Figure 4.4A. Mass spectrum of heme – dihydroartemisinin adduct synthesized in vitro. Adduct synthesized using ferriprotoporphyrin IX heme, glutathione, dihydroartemisinin, and excess phosphatidyl choline. Adduct was extracted from solution using 2% SDS and 5% guanidinium-Cl to mimic the FPIX extraction process used for live parasites (see Methods). Reproduced from [Heller et al., 2018b] without permission.

Our perfected FPIX heme extraction protocol [Heller et al., 2018a] entails treatment with excess sodium dodecyl sulfate (SDS) followed by precipitation of the detergent with guanidinium hydrochloride. In our hands, these treatments result in loss of H+ and addition of one Na+ (coordinated to tetrapyrrole propionate) to yield the observed ion at m/z 881 (Figure

4.4B).

85 N N N N N N Fe DHA Fe H +Na Fe H N N N N HO -H N N HO HO HO H H O O OH O O OH O OH O OH OH O OH OH O ONa m/z = 616 m/z = 858 m/z = 881 2+ 2+ FPIX (Fe , "B") FPIX (Fe )-C4-DHA, ("D")

Figure 4.4B. Mass spectrometry ionization of heme – dihydroartemisinin adduct synthesized in vitro. Fragmentation yields m/z = 881. Reproduced from [Heller et al., 2018b] without permission.

When FPIX is reacted in vitro with artesunate (ATS) instead of DHA (mass of 384 Da for

ATS vs 284 Da for DHA), the m/ z 881 peak shifts to m/z 981 (data not shown), which is consistent with this interpretation. When FPIX is not reduced with glutathione prior to addition of DHA, no adduct is formed (Figure 4.4C).

100 250

557.1

200 50

616.2 Intensity (kCounts) Intensity 150

0 810 830 850 870 890 910 m/z

100 498.1 Intensity (kCounts) Intensity

0 410 450 490 530 570 610 650 690 730 770 810 850 890 930 m/z

Figure 4.4C. Mass spectrum of the reaction between heme and dihydroartemisinin without addition of glutathione. No adduct 881 m/z is observed (inset=adduct region). Reproduced from [Heller et al., 2018b] without permission.

Following a similar approach as in the previous paper in this issue [Heller et al., 2018a] for FPIX quantification, quantification of FPIX−DHA LC−MS data is shown in Figure 4.4D and yields a linear relationship similar to that found previously for underivatized FPIX heme.

2500

2000

1500

1000

500 Signal Intensity (kCounts) Intensity Signal

0 0 500 1000 1500 2000 [Adduct] (nM)

Figure 4.4D Mass spectrometry calibration curve of ferriprotoporphyrin IX – dihydroartemisinin adduct. R2 = 0.90. Reproduced from [Heller et al., 2018b] without permission.

We next tested for the presence of FPIX−DHA adducts in live malarial parasites treated with bolus dose DHA for 6 h (see Methods). As shown in Figure 4.4.A, using essentially the same extraction procedure as that used for FPIX [Heller et al., 2018a], adduct is easily found for trophozoite stage CamWT parasites (Figure 4.5A; note clear peak at 881 m/z characteristic of adduct extracted under these conditions (Figure 4.5A inset; compare to Figure 4.4A).

Figure 4.5A. Adduct isolated from parasites after bolus dosing parasites with 5 µM + dihydroartemisinin for 6 hours. Heme species are seen at m/z 616.3 (M ), 557.2 (M-C2H3O2), and 659.4 (M-2H +2Na) [Robert et al., 2002a; Pashynska et al., 2004]. Phospholipid species is seen at m/z 760.6, and FPIX-DHA adduct is seen at m/z 881.3 (compare to Fig. 3A). Inset adjusts LCMS range to the narrower window of m/z 800-920 corresponding to the adduct region. Parameters for inset data collection were adjusted for visual representation to optimize the signal at m/z 881, with capillary voltage increased to 147.0 Volts and radio frequency loading decreased to 105%. Reproduced from [Heller et al., 2018b] without permission.

Importantly, the efficiency of CHCl3 extraction of the FPIX−DHA adduct is similar to the efficiency of extraction of free FPIX [Heller et al., 2018a] and is near 100% (Figure 4.5B).

88 1150

950

750

550

350 Signal Intensity (kCounts) Intensity Signal

150

-50 0 100 200 300 400 500 600 700 800 [Analyte] (nM)

Figure 4.5B. Calibration curves for known amounts of ferriprotoporphyrin IX heme and ferriprotoporphyrin IX – dihydroartemisinin adduct after extraction show similar extraction efficiencies. Squares: heme, Triangles: heme – dihydroartemisinin adduct. Extraction involved sodium dodecyl sulfate, guanidinium-hydrochloride and chloroform as described, followed by drying and re dissolving in methanol. The data show that the extraction procedure used in this work extracts known amounts of FPIX and FPIX-DHA adduct to near completion and with equal efficiency. Reproduced from [Heller et al., 2018b] without permission.

The FPIX−DHA adduct is also easily extracted for ring and schizont stage CamWT parasites (Figure 4.6A,E, left-hand side) treated with DHA in the same manner; however, absolute amounts are smaller presumably because of the lower abundance of free FPIX at these stages (Chapter 3) [Heller et al., 2018a]. Because of the altered abundance of free FPIX found for DCP parasites relative to the control (Chapter 3) [Heller et al., 2018a], we synchronized DCP parasites and extracted the FPIX−DHA adduct using identical methods. As expected, because the

FPIX abundance is significantly decreased at the trophozoite stage for Cam580Y (C580Y) relative to CamWT (Chapter 3) [Heller et al., 2018a], significantly less FPIX−DHA adduct is found at the trophozoite stage for Cam580Y treated identically to CamWT (Figure 4.6D vs

Figure 4.6C, and Figure 4.7) However, when the adduct is extracted from identically treated ring or schizont stage parasites, surprisingly, less adduct is again found for CamC580Y DCP parasites

89 relative to the CamWT control (Figure 4.6B vs Figure 4.6A, Figure 4.6F vs Figure 4.6E, and

Figure 4.7) even though the DCP parasites show slightly elevated levels of free FPIX at these stages (Chapter 3) [Heller et al., 2018a].

90 60 60 A B 50 50

40 40

30 30

20 20 Intensity (kCounts) * Intensity (kCounts) 10 10 *

0 0 830 850 870 890 910 830 850 870 890 910 m/z m/z

60 60 C D 50 * 50

40 40

30 30

20 20 Intensity (kCounts) Intensity (kCounts) * 10 10

0 0 830 850 870 890 910 830 850 870 890 910 m/z m/z

60 60 E F 50 50

40 * 40

30 30

20 20 Intensity (kCounts) Intensity (kCounts) * 10 10

0 0 830 850 870 890 910 830 850 870 890 910 m/z m/z CamWT CamWT-C580Y

Figure 4.6. Ferriprotoporphyrin IX – dihydroartemisinin adducts measured by mass spectrometry for mid-ring (A,B), trophozoite (C,D), and schizont (E,F) stages of control CamWT (left) and delayed clearance phenotype CamWT-C580Y (right) parasites. Asterisks indicates position of m/z 881 FPIX-DHA adduct peak. Reproduced from [Heller et al., 2018b] without permission.

91 30

25 M) µ 20

10 [Adduct in Parasite] ( Parasite] in [Adduct

0 CamWT Cam3.11-Revertant CamWT-C580Y Cam3.11-R539T

Figure 4.7. Quantification of ferriprotoporphyrin IX – dihydroartemisinin adducts for control CamWT and Cam3.11-R539T-Revertant strains versus delayed clearance phenotype CamWT-C580Y and Cam.311-R539T parasites. Black bars: mid-rings, Blue bars: mid-trophozoites, green bars: mid-schizonts. Data (+/- S.E.M.) represent at least two independent experiments (two independent parasite culture growths) with two replicates (two independent adduct measurements) of each (4 determinations in total). All stage - dependent differences between either delayed clearance phenotype parasite and its matched isogenic control are statistically significant according to Students t-test (p < 0.05). Reproduced from [Heller et al., 2018b] without permission.

To test the association between altered FPIX−DHA adduct levels and the DCP, a second

DCP strain, Cam3.11 R539T, and its isogenic control strain that does not harbor PfK13 mutations (strain Cam 3.11 R539Trev) [Straimer et al., 2015] were also analyzed. As shown in

Figure 4.7, although the absolute abundance differs somewhat relative to the control CamWT and DCP Cam580Y strains, a similar trend of reduced adduct abundance at all intraerythrocytic developmental stages is found for DCP strain Cam 3.11R539T relative to that of its genetically matched control, strain Cam3.11-R539T-Revertant. Upon quantifying these data versus those in

Chapter 3 [Heller et al., 2018a], we find that, depending on the strain, 0.3−0.5 and 0.6−1.2% of

92 total free FPIX present at the trophozoite and ring stages, respectively, are converted to the

FPIX−heme adduct.

Finally, to test whether the FPIX−DHA adduct is incorporated into or associated with hemozoin crystals, hemozoin from DHA-treated parasites was isolated and solubilized as described in Heller et al., 2018a. LC−MS data reveal only the FPIX monomer and reciprocal dimer for these samples (Figure 4.8) without measurable FPIX−DHA adduct or FPIX dimer−DHA adduct [note the lack of a peak at m/z 881 (Figure 4.8)].

700

616.5

600

500

400

300 Intensity (kCounts) Intensity

200 659.5 1231.9

1254.5 100

0 400 500 600 700 800 900 1000 1100 1200 1300

m/z

Figure 4.8. Mass spectrum of hemozoin isolated from CamWT parasites treated with dihydroartemisinin. Hemozoin was solubilized using NaOH [Trager et al., 1976]. Both FPIX monomer (616.4) and reciprocal dimer peaks at 1253.3 and 1231.3 [2*FPIX – 2H, + Na] and [2*FPIX -H], respectively are observed [Pashynska et al., 2004], but no FPIX-DHA adduct is observed (compare to Fig. 4A). Reproduced from [Heller et al., 2018b] without permission.

93 4.3 Discussion

The results of this study may be summarized as follows.

(1) A covalent FPIX−DHA adduct, similar to what is easily produced in vitro using mixtures of pure DHA and pure FPIX in the presence of an appropriate reducing agent, is also produced within live intraerythrocytic P. falciparum malarial parasites treated with plasma levels of DHA.

(2) The relative abundance of the FPIX−DHA adduct varies versus the stage of intraerythrocytic parasite development, with the largest abundance found at the trophozoite stage, corresponding to the stage that harbors the highest concentration of free FPIX heme within the parasite DV.

(3) The abundance of the FPIX−DHA adduct is lower for DCP parasites relative to matched isogenic control strains at all stages of parasite development.

(4) Although the FPIX−DHA adduct is formed within live parasites, no appreciable DHA or

FPIX−DHA adduct is found incorporated into hemozoin isolated from DHA-treated parasites.

It has been debated for some time whether FPIX heme is the primary activator of

Artemisinin prodrugs to produce Artemisinin drug radicals and whether covalent Artemisinin drug−FPIX heme adducts are formed within P. falciparum malaria. This study provides important data on both points and is the first to the best of our knowledge to directly measure the

FPIX−DHA adduct for live P. falciparum. One previous study reported the presence of the heme− Art adduct in the bloodstream of mice infected with P. vinckei vinckei malaria and subsequently treated with Art [Robert et al., 2005], but the origin of heme within the adduct and the site of adduct formation were not determined. Here we show that adduct FPIX is clearly formed within the P. falciparum parasite and that the adduct is likely formed within the parasite

DV.

The FPIX−DHA adduct is formed within live parasites treated with clinically relevant

94 plasma levels of DHA (these initial experiments are performed at 5.0 µM DHA for 6 h; the plasma DHA concentration ranges from 0.5 to 5 µM and perhaps even higher [Newton et al.,

2002; Khanh et al., 1999]). This observation has very important implications for the mechanism of action of Art drugs. FPIX−DHA adducts are likely quite toxic [Loup et al., 2007], with toxicity similar to that of FPIX alone, and perhaps even more toxic because the DHA moiety attached to FPIX would be predicted to improve membrane permeability properties allowing wider organellar access for the FPIX−DHA adduct relative to underivatized FPIX.

We find that the FPIX−DHA adduct abundance is lower for two DCP parasites expressing two different DCP-associated mutant PfK13 proteins versus their matched isogenic control parent strains, at all stages of parasite development. The observation of a lower adduct for DCP trophozoites versus control trophozoites is not entirely surprising because DCP trophozoites harbor reduced levels of free FPIX relative to control trophozoites (Chapter 3)

[Heller et al., 2018a]. However, the observation of reduced adduct for DCP rings and schizonts is curious because DCP parasites harbor slightly increased levels of free FPIX relative to control parasites at these stages (Chapter 3) [Heller et al., 2018a]. This point is discussed further below.

We propose that the lower abundance of the ring stage FPIX−DHA adduct for DCP parasites is a key feature of the molecular mechanism of evolving artemisinin resistance. Importantly, lower abundance is found for two different DCP strains harboring two different DCP-associated PfK13 mutations, suggesting that lower adduct abundance is a general feature of PfK13-mediated DCP.

However, for lower FPIX−DHA adduct to be correlated with DCP, two riddles must be explained. First, the toxic FPIX−DHA adduct is more abundant for trophozoites, yet several studies have found that Art drugs are more potent versus rings relative to trophozoites. Second, most studies have found that the reduced potency of Art drugs seen for DCP parasites is specific

95 to the ring stage [Witkowski et al., 2013; Amaratunga et al., 2014]. If the FPIX−DHA adduct is more abundant for trophozoites, how is ring stage Art drug selectivity explained? Obviously, much more work remains to be done related to this point, but we propose that the FPIX−DHA adduct targets metabolic processes specific to the ring more than metabolic processes specific to the trophozoite. Possibilities include dramatically upregulated transcription and translation that occurs at the ring stage, early ring stage Hemoglobin catabolism that may differ from trophozoite catabolism [Witkowski et al., 2013], ring stage membrane biogenesis, and/or organellar development. If the adduct affects these processes, then even small differences in the level of ring stage adduct abundance could conceivably confer DCP [Klonis et al., 2011].

It is well-known that DCP parasites do not show shifts in drug IC50 values similar to how chloroquine resistant (CQR) parasites show 10-fold shifts in chloroquine (CQ) IC50 values relative to control. This predicts that the decrease in adduct abundance for DCP trophozoites is not significant enough to have an equally profound effect on growth inhibition. With regard to this point, we note that adduct abundance corresponds to approximately 0.3−0.5% of the total amount of free FPIX present at the trophozoite stage (depending on the strain of the parasite); however, adduct abundance corresponds to 0.6−1.2% of the total amount of free FPIX present at the ring stage. We suggest that the greater percent abundance for rings is sufficient to promote ring stage quiescence as previously observed [Witkowski et al., 2010; Teuscher et al., 2012] but not to shift drug IC50 values.

With regard to the second riddle, the adduct concentration is found to be between 1 and

25 µM depending on the stage and strain; for CamWT trophozoites, it is ~20 µM. We propose that an adduct toxicity threshold is achieved for control strain rings and that this threshold is well surpassed for trophozoites. Via this reasoning, the decrease in trophozoite adduct abundance for

96 DCP parasites then, although significant, still leaves >5 µM levels of adduct present for DCP trophozoites, presumably still above this threshold, leading to only small effects on Art drug toxicity for trophozoites. In contrast, ~5 µM adduct is found for control strain ring stages, and

~2.5 µM adduct for DCP ring stages. We propose that the decrease in the DCP ring adduct concentration moves the level of FPIX−DHA adduct below the threshold relevant for toxic effects leading to quiescence, whereas copious remaining adduct in the DCP trophozoite or schizont is still sufficient to kill parasites at these stages. This idea predicts that the evolution of a heightened DCP correlated with further decreases in adduct abundance at these stages would promote formal ArtR.

Interestingly, we find a lower abundance of FPIX−DHA adduct for DCP rings and schizonts relative to control even though DCP rings and schizonts show increased levels of free

FPIX at these stages relative to the control. With regard to this apparent paradox, we note that the level of free FPIX is not necessarily the relevant moiety for activating Art -based drugs; rather, it is the level of ferrous FPIX (Fe2+FPIX) that is relevant. Ferrous FPIX is provided either upon immediate release of FPIX (as Fe2+FPIX) from catabolized Hemoglobin or upon reduction of the more abundant pool of FPIX within the parasite DV [likely ferric FPIX (Fe3+FPIX)]. The principal reductant that would control Fe2+:3+ ratios for any eukaryotic cell is glutathione.

Reduced glutathione is capable of donating e- to Fe3+ (Figure 4.2) and in the process is oxidized to GSSG (the GSH dimer). GSH:GSSG ratios are typically ≥20 for eukaryotic cytosol (the cytosol is highly reducing), whereas they are typically closer to 1 for lysosomes or vacuoles

[Noctor et al., 2013]. This predicts that the ratio of ferrous to ferric FPIX within the DV is low, yet ferrous FPIX is not zero. We estimate that the concentration of total FPIX is near 4 mM for control trophozoite DV (Chapter 3) [Heller et al., 2018a]. If even a modest 1% of the total FPIX

97 is reduced by available GSH, this predicts 40 µM ferrous FPIX heme would be available for activating DHA. In vitro, we and others [Hassett et al., 2017b; Robert et al., 1997] find that

DHA is activated to near completion in the presence of ferrous FPIX. Therefore, micromolar levels of ferrous FPIX within the DV are more than adequate for generating the levels of adduct that we observe. Lower levels of adduct for DCP rings, in spite of a higher total free FPIX concentration relative to the control, suggests a change in either the release of ferrous FPIX from catabolized Hemoglobin or GSH/GSSG traffic near the site of free FPIX (presumably, the nascent ring DV). Indeed, other work has found altered GSH/GSSG cycling linked to Art-based drug resistance phenomena [Rocamora et al., 2018; Dwivedi et al., 2017].

We suggest that once the DHA endoperoxide is activated (by Fe2+FPIX), its proximity to

FPIX, stabilized somewhat by an Fe−oxo radical bridge (Figure 4.2B), confers “preferred target” status on free FPIX, leading to efficient adduct formation within P. falciparum as observed.

However, as shown previously (Chapter 3) [Heller et al., 2018a; Wang et al., 2015]. other targets for DHA adduct formation are clearly possible within the parasite, but the percent of these targets that are effectively alkylated or how they would exert toxic effects sufficient to kill the parasite quickly is currently not known. Much work clearly remains to be done to define the rank order of DHA “preferred targets”. Here, the FPIX− DHA adduct is found to be an abundant intracellular poison for DHA-treated P. falciparum whose level is correlated with DHA sensitivity in DCP versus matched isogenic control strains.

To the best of our ability to ascertain, all known DCP parasites evolve within a chloroquine resistant (CQR) background. That is, although ArtR can be selected in the laboratory via other pathways, to the best of our knowledge, all naturally occurring DCP parasites that have been studied to date are also CQR. Association between CQR and DCP status could simply be a

98 consequence of the fact that most parasites found in southeast Asia (SEA), where DCP largely originates, are CQR, lowering the statistical likelihood of finding CQS/ DCP parasites. However, the CQR phenotype is known to be characterized by the presence of one of many different possible CQR-associated mutant PfCRT proteins [Callaghan et al., 2015], as well altered digestive vacuolar (DV) physiology, including changes in DV volume and pH [Eastman et al.,

2016; Bennett et al., 2004; Gligorijevic et al., 2006a; Gligorijevic et al., 2006b]. Whether CQR- associated alterations in DV physiology are obligate, permissive, or merely coincidental for the development of DCP parasites is not known; however, interestingly, some studies have suggested that some PfCRT mutations may in some way facilitate PfK13-mediated DCP

[Eastman et al., 2016]. Along these lines, CQR parasites are measured to have lower DV pH and other changes in DV physiology [Bennett et al., 2004; Gligorijevic et al., 2006a; Gligorijevic et al., 2006b; Roepe 2011; Lewis et al., 2014; Dzekunov et al., 2000], which are predicted to alter the availability of soluble FPIX. Changes in DV physiology measured for the CQR parasite DV might contribute to DCP via affects on FPIX−DHA solution chemistry.

In addition, studies have suggested that PfCRT and a PfCRT plant ortholog may transport

GSH [Patzewitz et al., 2000; Maughan et al., 2010]. If so, perhaps some mutant isoforms of

PfCRT associated with SEA CQR phenotypes facilitate important alterations in GSH/GSSG traffic that help to bias ferrous:ferric FPIX ratios. Lower ratios would act to decrease adduct abundance by lowering the Fe2+FPIX concentration necessary for Art-based drug activation and thereby facilitate the emergence of DCP. We note that a previous study has suggested higher levels of GSH in some CQR parasites; however, no direct measurements of GSH were taken in this work, and subcellular localization of any increased GSH was not determined [Meierjohann et al., 2002].

99 Finally, we note that, formally, there are two possible sources for reduced FPIX: either

DV localized or DV in origin but cytosolically localized after diffusion of DV FPIX. The cytosol is a highly reducing environment that would strongly bias FPIX toward the ferrous form.

Formally then, to explain lower adduct abundance, DCP parasites might harbor decreased concentrations of DV GSH or show a reduced level of transport of free FPIX from the DV to the cytosol. Relevant to this point, a recent study finds ~1.5 µM heme within the cytosol that does not change across the parasite life cycle [Abshire et al., 2017], suggesting that the large differences in free FPIX and FPIX− DHA adduct that we observe across life cycle stages are clearly due to DV-localized FPIX.

100 1 CHAPTER 5. ARTESUNATE ACTIVATION BY HEME IN AN AQUEOUS MEDIUM

5.1 Introduction

Malaria remains a serious challenge to human health. Annually this disease affects about three billion people living in malaria-endemic regions as well as international travelers, governmental workers and deployed military personnel. Previous first - line drugs such as chloroquine (CQ) and sulfadoxine-pyrimethamine (SP) are no longer viable in many regions due to widespread drug resistance. These drugs have largely been replaced by artemisinin combination therapies (ACTs) composed of a derivative of artemisinin and one of several possible "partner" drugs (Figure 5.1) [Tu 2011].

Figure 5.1. Structures of antimalarial drugs used in this analytical study. Drugs denoted as ATM, ATS and DHA are derivatives of the parent drug artemisinin that are commonly used in the malaria clinic. Additional drugs LUM, PPQ, and AQ are commonly used "partner" drugs in artemisinin combination therapies (ACTs).

1Submitted to Inorganic Chemistry in April 2019 by Laura E. Heller, Paul D. Roepe*, and Angel C. de Dios*

101 Unfortunately, recently, a harbinger of artemisinin resistance known as the "delayed clearance phenotype" (DCP) has been reported across Southeast Asia [Ashley et al., 2014].

Consequently, dihydroartemisinin (DHA) with piperaquine (PPQ; Figure 5.1) as a partner drug, followed by a single low-dose primaquine (PQ) treatment is now recommended in areas of

Cambodia where both artemisinin resistance (ArtR) and PPQR is emerging [Amaratunga et al.,

2016; Dondorp et al., 2009]. The molecular pharmacology of these drugs remains incompletely understood. Elucidating this pharmacology is essential for understanding pathways to resistance and for the development of next generation therapies.

The Art 1,2,4 trioxane, specifically its bridging endoperoxide group, has been shown to be crucial for drug activity [Asawamahasakda et al., 1994; Robert et al., 2005; Meunier et al.,

2010; O’Neill et al., 2010; Hassett et al., 2017b]. However, the detailed molecular mechanism of endoperoxide activation within the malarial parasite, and how the activated drug then kills the parasite remain active areas of research. The trioxane group is highly reactive [Herwig et al.,

2007], and pioneering work of Meshnick et al. [Meshnick et al., 1991] suggested that antiparasitic reactivity requires activation by Fe(II). Two competing models have been proposed to describe this reaction. One envisions initial Fe(II) mediated cleavage of the peroxide bond

[Posner et al., 1992; Posner et al., 1994; Posner et al., 1995; Jefford et al., 1995; Jefford et al.,

1996]. A second model involves ring opening driven by either protonation of the peroxide or action by a Lewis acid [Haynes et al., 1996a; Haynes et al., 1996b; Haynes et al., 1999; Haynes et al., 2007], thereby assigning only a minor role to Fe(II). Recent work suggests that heme

Fe(II), and not necessarily labile Fe(II), may account for the activation of Art - based drugs within malarial parasites [Wang et al., 2015; Heller et al., 2018a; Heller et al., 2018b]. Two studies have shown that > 100 parasite proteins are potential Art drug alkylation targets [Wang et

102 al., 2015; Heller et al., 2018a; Heller et al., 2018b; Ismail et al., 2016a], but overlap between these two proteomic data sets is limited, suggesting protein alkylation by activated Art - based drugs may be somewhat random [Hassett et al., 2017b; Heller et al., 2018b]. One recent study has found that activated DHA alkylates free FPIX within parasites [Heller et al., 2018b] and synthetic endoperoxide - based antimalarials have been shown to alkylate proteins [Ismail et al.,

2016b]. Currently it is unknown which Art - based drug adducts produced within the parasite are pharmacologically the most relevant. DHA - FPIX adducts are known to be toxic and some protein adducts might be predicted to activate stress response and /or cell death pathways, suggesting that both protein and FPIX adducts may be relevant.

The promiscuous targeting of Art and its activation by FPIX heme are somewhat consistent with current data on emerging Art resistance (ArtR), also known as the "delayed clearance phenotype" (DCP). Specifically, PfKelch13 amino acid substitutions are associated with DCP/ArtR primarily at the ring stage of parasite development [Witkowski et al., 2013;

Ariey et al., 2014; Straimer et al., 2015] during which Art is not as efficiently activated due to lower availability of free FPIX relative to other parasite stages [Heller et al., 2018a; Heller et al.,

2018b]. It has been suggested that increasing Art treatment duration [Dogovski et al., 2015] or the use of δ-aminolevulinic acid, a known heme precursor, may be effective in combating ArtR

[Wang et al., 2015].

FPIX heme is liberated during obligate parasite hemoglobin (Hb) catabolism that occurs during parasite red blood cell development, and is also a known target for other antimalarial drugs. For example, quinoline-based drugs such as chloroquine (CQ), quinine (QN) and amodiaquine (AQ) interact directly with FPIX, and prevent its crystallization to inert hemozoin

[Leed et al., 2002; Casabianca et al., 2008; Gorka et al., 2013c; de Dios et al., 2004]. Using

103 nuclear magnetic resonance (NMR) spectroscopy and the effects of the FPIX paramagnetic

Fe(III) center on the relaxation rates of drug protons, solution structures of noncovalent complexes formed between quinoline - based drugs and FPIX have been determined [Leed et al.,

2002; de Dios et al., 2004]. Furthermore, it is now known that an equilibrium exists between monomeric and dimeric FPIX species. This equilibrium has been extensively studied as a function of pH, and in the presence of either CQ or QN [Casabianca et al., 2008]. At low pH, the monomeric species is favored while at higher pH values, the antiferromagnetic dimer dominates. The transition in purely aqueous medium occurs at a pH lower than 5; only the antiferromagnetic species is then observed as precipitation begins to occur upon protonation of the heme propionic acid side chains. Abundant monomeric species is therefore observed only if a mixture of DMSO and H2O is used as solvent. In these previous studies, CQ was found to promote dimer formation while QN promotes the converse, highlighting that even closely related quinoline antimalarial drugs may have different molecular mechanisms of action [Gorka et al.,

2013c]. The state of FPIX in solution is clearly consequential to how it interacts with quinoline antimalarial drugs,30-33 [Leed et al., 2002; Casabianca et al., 2008; Gorka et al., 2013c; de Dios et al., 2004] and is likely also important for interaction with Art - based drugs, which are activated by Fe(II)FPIX [Robert et al., 2005; Meunier et al., 2010; Meshnick et al., 1991; Wang et al., 2015; Heller et al., 2018b]. These issues are critical for elucidating the pharmacology of some ACTs, which combine Art - based and quinoline drugs.

Art - based drugs act rapidly against P. falciparum, and are eliminated quite rapidly, having a half-life in plasma of about 1 to 2 hours. Effective ACTs thus include drugs with longer half-lives such as LF, PPQ, and AQ (Figure 5.1) [Eastman et al., 2009; Henrich et al., 2014].

When FPIX is released upon parasite Hemoglobin digestion, Fe(II) is presumably oxidized to

104 Fe(III) due to the presence of molecular oxygen. For Art activation, Fe(II) is required. Therefore, if Fe(II)FPIX is relevant for Art activation within malarial parasites, the drug must either encounter FPIX as it is released during Hemoglobin catabolism or a reducing agent (presumably

GSH) must convert Fe(III) to Fe(II). GSH is found at significant concentrations inside the parasite [Atamna et al., 1997]. Using GSH to reduce Fe(III)FPIX, efficient DHA alkylation of heme has previously been demonstrated [Robert et al., 2002a; Heller et al., 2018b].

Unfortunately, most in vitro studies involving FPIX and Art - based drugs have been performed in organic solvents or mixed aqueous-organic medium that reduce FPIX self-association relative to an aqueous environment. Rapid FPIX self-association in aqueous environment can involve either cofacial π−π interactions or hydroxo-bridge bonds [Monti et al., 1999]. The formation of

Fe(II)/Fe(III) protoporphyrin IX complexes, on the other hand, has not been observed in DMSO

[Robert et al., 2002a], indicating that the behavior of FPIX, Art and GSH in an aqueous environment may be significantly different from what is observed in DMSO [Haynes et al.,

2005].

Notwithstanding these complex studies using aqueous - DMSO solutions have been, and remain, useful [Casabianca et al., 2008]. The higher viscosity relative to aqueous media dictates that FPIX protons can be readily observed by NMR spectroscopy due to favorable rotational correlation times. In acidic DMSO, the FPIX monomer is dominant resulting in NMR signals that display substantial paramagnetic shifts. The four peaks observed with chemical shifts greater than 55 ppm from tetramethylsilane (TMS) arise from the protons of the four inequivalent methyl groups attached to the porphyrin ring in FPIX. In theory then, observing the reaction between Art and FPIX in the presence of GSH in acidic aqueous DMSO solution would provide concrete evidence of alkylation of the porphyrin ring. Also, the 55-70 ppm chemical shift region

105 allows for non-invasive direct measurement of the rate of Fe(II)FPIX reduction by GSH. FPIX methyl peaks would disappear as Fe(III) is reduced to Fe(II), which would then correlate with appearance of oxidized glutathione (GSSG) peaks. As Fe(II) is oxidized back to Fe(III) upon reductive activation of the Art peroxide, methyl peaks would reappear. With subsequent alkylation of the FPIX porphyrin ring by Art, new methyl resonances would then be observed. In such studies, FPIX protons would not be directly observable, however, drug protons can be monitored.

We have used these principles to examine the kinetics of Art - based drug / FPIX interactions and how they are influenced by well-known FPIX aggregation. In addition to elucidating important factors that affect how Art is activated by FPIX, these data provide insight on how ACT partner drug interactions with FPIX may influence Art - derived drug potency.

5.2 Results

The reaction between FPIX and ATS was monitored by NMR spectroscopy in aqueous /

DMSO solution. Most ATS peaks in the 1D proton NMR spectrum are multiplets except for those due to protons of the methyl group attached to Carbon-3 and the proton attached to

Carbon-12 (Figure 5.2).

Figure 5.2. Artesunate (ATS) and the numbering scheme employed in this work.

106 In the absence of GSH and FPIX, the 3-CH3 signal, being a singlet and arising from three

1H, is the most intense in the ATS 1D spectrum (Figure 5.3).

Figure 5.3. The 3-methyl resonance of artesunate before (a) and after (b) reaction with reduced heme formed in situ from Fe(III) heme and glutathione. Peaks marked with "*" are tentatively assigned to acetate and acetyl proton.

This methyl is also proximal to the ATS endoperoxide (Figure 5.2) such that the 3-CH3 signal disappears upon ATS reaction with Fe(II)-protoporphyrin IX (Figure 5.3; compare b vs. a). The disappearance of 3-CH3 signal is quite dramatic and is accompanied by the appearance of a resonance near 2.0 ppm (Figure 5.3b), which can be assigned to the protons of an acetate ion or an acetyl group liberated during the reaction between ATS and Fe(II)FPIX [Posner et al., 1994;

Posner et al., 1995]. We studied the Fe(II)FPIX - ATS reaction in aqueous medium in two different ways. The first initially incubated FPIX with GSH to reduce all Fe(III) to Fe(II) before adding ATS. The second mixed aqueous solutions of FPIX, GSH and ATS simultaneously, such that ATS reacted with FPIX Fe immediately upon reduction of Fe. Overnight incubation of FPIX with GSH showed a significant decrease in the rate of FPIX Fe activation of ATS (once ATS

107 was added) as monitored by loss of the 3-CH3 resonance (Figure 5.4b), relative to the rate observed when FPIX, GSH and ATS were mixed simultaneously (Figure 5.4a).

Figure 5.4. NMR resonance signal of 3-methyl of artesunate obtained in 30 minute intervals from an aqueous sample containing ferriprotoporphyrin IX heme, glutathione and artesunate. (a) all materials mixed at beginning of experiment. (b) with overnight incubation of FPIX with GSH before adding ATS.

Using peak integrals to quantify loss of the 3-methyl signal, we compared these two reactions (Figure 5.5a). Here, the faster aqueous reaction has not reached completion even after 3 hr.

108

Figure 5.5A. Conversion of artesunate to C4 centered radical artesunate versus time. Experiments are with (top, ×) and without (bottom, ¿) prior complete reduction of heme by glutathione.

1 0.9 0.8 0.7 0.6 0.5 0.4

NMR intensity 0.3 0.2 0.1 0 Time, minutes 0 50 100 150 200 250 300 350 400

Figure 5.5B. Conversion of artesunate to C4 centered radical artesunate versus time (top, ¿) and oxidation of reduced glutathione (GSH) into oxidixed glutathione (GSSG) versus time (×).

When incubated alone with FPIX, neither GSH nor ATS bind as strongly to FPIX Fe(II) as do quinoline antimalarial drugs such as CQ [Leed et al., 2002]. NMR relaxation

109 measurements of ATS or GSH protons in the presence of FPIX do not show an appreciable decrease in longitudinal relaxation times (T1). The fact that NMR signals from ATS and GSH can still be observed in these samples suggests that the dissociation constant between either ATS

1 or GSH with FPIX is > 1 mM H attached to Carbon-10 or -12 in ATS demonstrate a T1 of about

0.8 s in D2O (not shown, see [31]). In the presence of Fe(III)FPIX in D2O, the T1 for these protons is reduced to only 0.6 s. In DMSO, the reduction in T1 is stronger; to 0.3 s, indicating a slightly higher affinity for FPIX monomer in this solvent. In contrast, proton resonances from

CQ are barely observed in aqueous samples that contain 1 mM CQ and 100 µM FPIX

[Casabianca et al., 2008].

As ATS reacts with reduced Fe(II)FPIX, new signals are observed near the methyl region of the proton NMR spectrum (Figure 5.3b). These new signals (asterisks, Figure 5.3b) are attributed to either an acetate or acetyl group (AcO) liberated during ATS alkylation of

Fe(III)FPIX (Figure 5.6) (Chapter 4) [Heller et al., 2018b].

Figure 5.6. Proposed mechanism for alkylation of ferriprotoporphyrin IX heme by artesunate. The growth of these signals does not occur at the same rate as the disappearance of the

ATS 3-CH3 resonance due to activation of ATS (Figure 5.3a). This is consistent with the formation of one or more alkylation intermediates as previously suggested41 and recently observed (Chapter 4) [Heller et al., 2018b].

110 Quinoline antimalarial drugs such as CQ interact strongly with FPIX in aqueous solutions

[Leed et al., 2002]. NMR studies in solution indicate that while interaction between quinoline drug and FPIX are significant, the dominant complex formed is noncovalent and there is fast exchange between free and bound drug [Leed et al., 2002; de Dios et al., 2004]. Nevertheless, when quinoline drugs bind to FPIX, access to the Fe center is hindered [Leed et al., 2002; de

Dios et al., 2004]. We therefore hypothesized that the reaction between Fe(II)FPIX and ATS might be negatively impacted by the presence of a quinoline drug. Using NMR experiments similar to those described above, (e.g., simultaneous mixing of aqueous solutions of FPIX at pH

7.0, GSH, ATS, and equimolar heme-targeted antimalarial (HTA) drug), we observed results similar to those obtained with prior incubation of FPIX with GSH (Figure 5.7).

Figure 5.7. The amount of unreacted artesunate as a function of time in the absence () vs presence of other FPIX - binding antimalarial drugs. Chloroquine (CQ, ¯), piperaquine (PPQ, ¿), amodiaquine (AQ, £), pyronaridine (PY, p), lumefantrine (LUM, +), quinine (QN, r), primaquine (PQ, !), and 9-epi-quinine (EPI, ). Namely, the reaction between Fe(II)FPIX and ATS is significantly impeded by the presence of CQ and ACT partner drugs such as AQ, but not by 9-epi-quinine (EPI), which binds

111 much more weakly to FPIX, in edge - to - face fashion [Gorka et al., 2013c]. Figure 5.8 shows that even with a CQ:FPIX molar ratio of 1:4, the rate of disappearance of 3-CH3 signal is still significantly slowed. With only one bound CQ molecule per FPIX [Leed et al., 2002], predicted

ATS activated is only 20 percent compared to control (compare Figures 5.8 vs. 5.5).

Figure 5.8. The amount of unreacted artesunate as a function of time in the presence of increasing ratios of chloroquine. Chloroquine:Heme ratios: 1:1 (), 1:2 (), 1:4 (£), and 0:1(!). Similar results are obtained with QN and AQ where only 30 and 20 percent of ATS is activated after 4 hours with drug:FPIX ratio of 1:4 (not shown). Correspondingly, in the presence of quinoline drug, the increase in GSSG peaks, specifically β-cysteinyl proton resonances, are significantly delayed (not shown). The quinoline drug therefore prevents reduction of

Fe(III)FPIX by GSH. Without reduced heme, ATS cannot be activated and FPIX cannot be alkylated.

Earlier results suggested that the diastereomer of quinine, 9-epi-quinine (EPI), is unique with regard to FPIX interaction relative to other quinoline drugs. Specifically, while CQ and QN

112 have been shown to bind to the face of the FPIX porphyrin plane [Gorka et al., 2013c], ring- current shifts measured with a diamagnetic analog ZnPIX suggested that EPI binds with a face- to-edge π- π interaction in which the quinoline ring is oriented along the side of the porphyrin

[Gorka et al., 2013a]. Edge binding would be predicted to stabilize but not cap FPIX aggregates, and would not therefore prevent ATS access to Fe(II) within FPIX. Initially, the rate of ATS activation is limited by the amount of exposed heme. However, as ATS reacts and alkylates

FPIX and frees a porphyrin unit from the aggregate, a new heme unit is now exposed. Thus, compared to the reaction when EPI is absent, the reaction is closer to being zero order as the concentration of heme available for reduction by GSH is nearly constant throughout the reaction.

Correspondingly, (Figure 5.7), the apparent initial rate of ATS activation is slower in the presence of EPI since with aggregation, the effective amount of FPIX available for reduction of

GSH is reduced.

FPIX - ATS and FPIX - ATM adducts were analyzed by ESI - MS and showed m/z of

941, 854, respectively (Table 5.1), as previously reported (Chapter 4) [Heller et al., 2018b].

When ATM is reacted in the presence of EPI, ESI - MS reveals the presence of a single FPIX molecule being alkylated twice (Table 5.1, Figure 5.9). A clear peak at m/z = 1092 (Figure 5.9a) can be assigned to a 1:2 FPIX:ATM adduct with both ATM moieties DE acetylated and one of the propionates bearing a Na+ ion (Figure 5.9b).

113

Figure 5.9A. Proposed structures for ferriprotoporphyrin IX heme - artemether adducts formed. Masses are 854 and 1092 in the presence of Epi quinine.

Figure 5.9B. Mass spectrometry peaks observed for ferriprotoporphyrin IX heme - artemether adduct formation in the presence of Epi Quinine. Note peaks at 854 and 1092 corresponding to left and right (see Figure 5.9a).

114 In the absence of EPI, only the monoalkylated adduct, m/z = 854, is observed (Table 1).

Double alkylation of FPIX in the presence of EPI suggests that an alkylated FPIX with a regenerated Fe(III) center can be reduced and then activate another ATM molecule. This is more likely to happen if the FPIX aggregate is stable (as when EPI is binding edge - to - face to FPIX units in the aggregate) and a singly-alkylated heme at the end of an aggregate remains the only

Fe(III) accessible to GSH.

Table 5.1. Observed m/z peaks for FPIX-ATM adduct formation in the presence of EPI, detected via electrospray ionization (ESI) mass spectrometry Sample Observed m/z peaks FPIX:GSH:ATM 854 FPIX:GSH:ATM:EPI 854, 1092 FPIX:GSH:ATS 941 FPIX:GSH:ATS:EPI 941, 1264

5.3 Discussion

Monitoring the rate of the loss of 3-CH3 signal provides a convenient method for measuring the rate of the reaction between Fe(II)FPIX and ATS. In pure DMSO, where FPIX is first incubated with GSH overnight, the reaction is quite fast such that it is complete before an

NMR spectrum can be obtained. In water, on the other hand, the results are notably different.

Comparing mixed aqueous solutions of FPIX, GSH and ATS simultaneously vs. reducing FPIX with GSH before addition of ATS, the difference in rate is easily explained by self-association of

Fe(II)-FPIX, as previously observed [Monti et al., 1999]. That is, it is well known that

Fe(II)FPIX readily self associates via oxo-bridges and π−π interactions, which would significantly reduce ATS access to the FPIX Fe(II) center, thereby lowering ATS activation efficiency as observed (Figure 5.4b). Interaction between FPIX and ATS is much more efficient without prior complete reduction of FPIX by GSH; As shown in Figure 5.5a, the faster aqueous reaction has not reached completion even after 3 hr. This is in stark contrast with the fast reaction

115 observed in DMSO [Robert et al., 2002a]. In a nonpolar solvent that maintains FPIX in monomeric form, FPIX alkylation by ATS followed by FPIX demetallation is complete within a few minutes [Robert et al., 2002b]. Reactant concentrations in the current work are an order of magnitude lower than those used by Robert and colleagues [Robert et al., 2002a]. In D2O, where FPIX protons are completely invisible, the reduction of Fe(III) in FPIX can be followed by monitoring oxidized GSH and/or reduced GSSG resonances, specifically those from β- cysteinyl protons. The appearance of GSSG peaks correlates with the disappearance of the 3-CH3 signal (data not shown). We find that, in aqueous media, and at physiologically relevant concentrations, the reaction between ATS and FPIX strictly correlates with conversion of GSH to GSSG, suggesting that the reduction of FPIX Fe(III) by GSH is rate limiting (Figure 5.5b).

Interaction between FPIX, GSH and Art leads to the scission of Art’s endoperoxide bring and eventual formation of a covalent FPIX-Art adduct. Alkylation of FPIX by ATS (Figure 5.6) proceeds via two main steps. First, Fe(II) in reduced FPIX breaks the ATS endoperoxide bridge, and due to the formation of an O-centered radical and proximity of the paramagnetic Fe(III), proton resonances from ATS are significantly perturbed in the NMR spectrum as an iron-oxo intermediate forms (Figure 5.6, left). The O-centered radical then rearranges to form a primary carbon centered radical (Figure 5.6, middle). This radical then alkylates the porphyrin ring of

FPIX (right) which gives rise to an acetyl peak in the proton NMR spectrum (see Figure 5.3b)

Since the appearance of free acetyl signal is delayed, the release of this acetyl group does not occur immediately after FPIX reduction and ATS activation (Figure 5.6). This delay may likewise suggest that alkylation of the porphyrin ring does not occur instantly. Intermediates prior to FPIX alkylation are highly reactive as these contain either O- or C-centered radicals

(Chapter 4) [Heller et al., 2018b]. If FPIX alkylation does not occur instantly, these

116 intermediate(s) might then facilitate alkylation of other targets such as parasite proteins [Ismail et al., 2016a] after diffusion of the ATS radical. This covalent ATS mediated sequestration of

FPIX is distinctly different from noncovalent quinoline antimalarial drug sequestration [Loup et al., 2007].

Quinoline drugs effectively prevent the reduction of Fe(III)FPIX by GSH, as even with a

CQ:FPIX molar ratio of 1:4, the rate of disappearance of 3-CH3 signal is still significantly slowed. Why equimolar quinolinal drug is not required for this effect illustrates the nature of

FPIX heme in aqueous medium. Access to the Fe(III) center would be limited even with a substoichiometric amount of the quinoline drug if heme exists in an aggregated form as proposed

[Ursos et al., 2001]. In such circumstances, only the ends of the FPIX aggregates would provide

Fe(III) for GSH reduction. When a quinoline drug caps the end of these aggregates [Leed et al.,

2002], thereby denying access to GSH, Fe(II) for activating ATS is less available. These aggregates have been suggested to be composed of FPIX dimers or monomers interacting via

π−π interactions [de Villiers et al., 2007]. These aggregates are distinct from crystalline hemozoin (Hz) [Pagola et al., 2000; Hempelmann et al., 2002]. Capping these aggregates presumably assists inhibition of hemozoin formation [Ziegler et al., 2001]. Previous NMR studies [Leeed et al., 2002] of CQ binding to FPIX in aqueous medium used large excess of CQ

(≥ 10 fold) relative to FPIX, which is sufficient to inhibit FPIX aggregation. However, without such a large excess of CQ, FPIX aggregates in aqueous medium even at neutral pH [de Villiers et al., 2007]. In the presence of CQ and other quinolinal antimalarial drugs that are positively charged at neutral pH, aggregation can be enhanced by balancing the negative charge accumulating within the aggregate. This then explains why a substantial excess of drug is required to prevent FPIX self-association.

117 In sum we find that reduced FPIX reacts with ATS to form FPIX - ATS alkylated species and that the rate is limited by the reduction of Fe(III) to Fe(II) in FPIX by GSH. This reaction is found to be dependent on the behavior of FPIX in aqueous solution. Aggregation and capping of the aggregates by other antimalarial drugs can slow or prevent activation of ATS by FPIX.

However, in the special case of EPI, which binds very differently to FPIX, aggregates are not capped but are instead stabilized by binding of EPI molecules on the sides of the aggregate. The

Fe(III) center at the ends of these aggregates remain accessible for reduction by GSH.

118 CHAPTER 6. SYNTHESIS AND ANALYSIS OF TWO ARTEMISININ DRUG PROBES: LOCALIZATION

AND COMPETITION ANALYZED VIA SPINNING DISK CONFOCAL MICROSCOPY

6.1 Introduction

Endoperoxide antimalarials such as artemisinins (Arts) are components of current first- line drug combination treatments known as artemisinin combination therapies (ACTs) for

Plasmodium falciparum malaria as recommended by WHO. The Art class of antimalarials is defined in part by a highly reactive endoperoxide bridge that is cleaved by Fe(II) to produce radicals which then alkylate numerous targets within malarial parasite (see Chapter 1 and

Chapter 4). The exact mechanism for activation of this artemisinin pharmacophore and its relation to parasite death remain an active area of research. Pioneering work [Meshnick et al.,

1991] involved extensive studies on Art – heme interactions, and showed that a heme Fe (II) can activate Art endoperoxide groups in vitro. Recently, mass spectrometry (MS) – based metabolomic analysis of live intraerythrocytic malarial parasites has provided a greater understanding of Plasmodial parasite metabolite levels and metabolic responses to several common antimalarial drugs [Creek et al., 2016; Llinas et al., 2016] Taking advantage of Art drug covalent interactions, MS – based metabolomics can also be used to define drug targets, including for Art drugs. MS – based drug target profiling for Art-sensitive (ArtS) vs. Art- resistant (ArtR) parasites should help elucidate mechanisms of ArtR and define key concepts for the development of next generation ACTs.

Two earlier proteomic studies used incorporation of a biotin moiety onto Art, for streptavidin affinity “pull down” of target proteins covalently attached to Art drug probes. For effective isolation of the probe-labeled proteins, the entire drug probe must be stable in culture medium.

This is not a trivial requirement as Art drugs are not only known to hydrolyze in solution, but

119 their endoperoxide group is extremely reactive and, once cleaved, can cause a breakdown of the drug into a non-reactive deoxy form of artemisinin. [Posner et al., 1992] The O’Neill group

[Barton et al., 2010] were the first to synthesize a biotinylated artemisinin probe with nanomolar in vitro activity vs several strains of P. falciparum, however results using the probe in proteomics studies were not published until 2016 [Ismail et al., 2016a].

The O’Neill group [Ismail et al., 2016a] and the Lin group [Wang et al., 2015] have each published on alkyne-tagged Art derivatives, both able to be coupled with a biotin azide (i.e.

“clickable” Art probes). Clickable alkyne probes such as these can be useful to isolate drug targets through a process called activity-based protein profiling (ABPP) [Raghavan et al., 2009;

Best 2009]. For ABPP, the probe must have one reactive subunit allowing it to attach to the target (i.e. the drug portion of the probe) and another subunit used for the detection or selective purification of probe-target complexes (Figure 6.1). In the case of Art drugs, the reactive subunit is the Art endoperoxide (Figure 6.2).

Figure 6.1. A drug probe derivatized with an alkynyl tag. Following covalent linkage to the drug’s target(s), the alkyne handle can be reacted with an azide labeled fluorophore or azido biotin, allowing for visualization or purification. Reproduced from [Best et al., 2009] with permission.

120 O O Receptor Bioorthogonal HO O binding HO labelling H3C O O H C O 3 H H O O O O N3

N N

Figure 6.2. An alkynyl artemisinin probe. Following linkage to artemisinin’s target(s), the alkyne handle can be reacted with an azide labeled fluorophore or azido biotin yielding artemisinin bound to both its target(s) and a fluorophore or azido biotin tail for further analysis.

In ABPP, labeling of drug targets is typically followed by proteome analysis, which can be facilitated using either gel-based or avidin-affinity techniques. Following separation of the targeted proteins, MS is used for identification of trypsin digestion products of target protein(s).

[Salisbury et al., 2007]

The two aforementioned proteomics studies [Ismail et al., 2016a; Wang et al., 2015] have shown that > 100 parasite proteins are potential Art alkylation targets, but overlap between these two proteomic data sets is limited, suggesting protein alkylation by activated Art - based drugs may be somewhat random (see Chapter 1, Section 1.5.5) [Hassett et al., 2017]. While eye- opening, these studies were both done using strain P. falciparum strain 3D7 parasites and therefore do not explore differences, if any, between ArtS vs. ArtR parasites. In theory there may be protein targets present in ArtS but not ArtR strains, or vice versa; additionally, the abundance of some may differ.

I have synthesized two Art derivatives, one a biotinylated ATS probe and one a clickable alkynyl Art that was then reacted with an azido fluorophore as well as a biotin azide. Both

121 probes’ subcellular localization was visualized using fluorescence confocal microscopy. The goal of synthesizing these two probes was to test for differences between ArtS and ArtR drug targets and their abundances. Two different probes were synthesized at the same time as it was unclear which type of probe would be more stable in culture and best mimic underivitized drug.

As larger chemical handles can in theory affect a probe’s ability to cross cell membranes, replacing the biotin tag with a small alkyne handle might allow this probe to diffuse into the cell more effectively and to better bind to targets.

6.2 Results

The synthesis of Biotinylated ATS (ATS-b) relied on amide coupling between a pentylamino biotin and the carboxylic acid Art derivative, artesunate (ATS) (1), Scheme 2.1).

This one pot reaction was straight forward and high yield after adding a phase transfer catalyst (t- butylammonium bromide), as otherwise ATS is only partially miscible in aqueous solution while the aminobiotin is water soluble. After mixing overnight at 50° C, the final product was isolated by organic extraction and purified with DCM to yield light tan crystals.

A second Art-drug probe (DHA-a) was synthesized which contained a clickable alkynyl tail.

This tail can be coupled to azide-tagged fluorophores or to biotin. As described (Section 2.2.8.2), this probe was synthesized directly from dihydroartemisinin (DHA), by acetylating DHA and replacing the acetyl group with a terminal alkyne ((2), Scheme 2.2).

ATS-b was first tested for its effectiveness against P. falciparum in an IC50 assay compared with underivatized ATS as a control. The probe largely retained its biological activity, as the IC50 values versus HB3 and Dd2 strains of P. falciparum were comparable to that of underivatized ATS (Figure 6.3, Table 6.1).

122 HB3 ATS IC50 Dd2 ATS IC50

120 120

100 100

80 80

60 60 % Growth % % Growth % 40 40

20 20

0 0

0.1 1 10 100 1000 0.01 0.1 1 10 100 1000 [ATS] (nM) [ATS] (nM) HB3 Probe IC50 Dd2 Probe IC50

120 120

100 100

80 80

60 60

% Growth % Growth % 40

20 20

0 0

-20 0.01 0.1 1 10 100 1000 0.01 0.1 1 10 100 1000 [Probe] (nM) [Probe] (nM) Figure 6.3. Biotinylated probe retains its antimalarial potency as quantified by IC50 assay, compared with underivatized artesunate against P. falciparum strains HB3 and Dd2.

Table 6.1. IC50 values for ATS and ATS-b probe. The IC50 of the probe is within four-fold that of the value for underivatized ATS. HB3 Dd2 ATS 7.25 ± 0.62 nM 4.23 ± 0.69 nM Biotinylated ATS Probe 26.06 ± 1.74 nM 18.44 ± 2.22 nM

In a manner similar to that for ATS-b, this alkynyl DHA probe was tested for its effectiveness against P. falciparum in a side by side IC50 assay vs. underivatized ATM (Figure 6.4). ATM was used as a control for its similar size as compared to the alkynyl probe. The probe was found to retain its biological activity, as IC50 values vs. HB3 and Dd2 were comparable to that of underivatized ATM (Table 6.2).

123 HB3 ATM IC50 Dd2 ATM IC50

120

100 100

80 80

60 60 % Growth %

% Growth % 40 40

20 20

0 0.1 1 10 100 10 100 [ATM] (nM) [ATM] (nM)

HB3 Alkyne IC50 Dd2 Alkyne IC50

120 120

100 100

80 80

60 60 % Growth % % Growth % 40 40

20 20

0 0

1 10 100 1 10 100 1000 [Alkyne] (nM) [Alkyne] (nM)

Figure 6.4. Alkynyl probe retains antimalarial potency via IC50 assay, relative to artemether against P. falciparum strains HB3 and Dd2.

Table 6.2. IC50 values for ATM and the alkynyl probe. The IC50 of the probe is within five-fold that of the value for underivatized ATM. HB3 Dd2 ATM 6.03 nM 16.17 nM Alkynyl Probe 25.52 nM 26.56 nM

After determining that the probes had retained biological activity, the subcellular localization of each probe was analyzed using spinning disk confocal microscopy. For analysis of the ATS-b probe parasites were incubated with ATS-b, fixed with 4% paraformaldehyde /0.004% glutaraldehyde, and permeabilized with Triton-X before incubation with AlexaFluor 647-

124 conjugated streptavidin (1:200 dilution) to detect the biotinylated drug probe. Figure 6.5 shows subcellular localization of ATS-b within mid-trophozoites and a side by side control experiment incubating trophozoites with AlexaFluor 647-conjugated streptavidin but no prior incubation with (non fluorescent) probe, suggesting that detectable fluorescence within the iRBC via the streptavidin fluorophore is only possible upon conjugation of the fluorophore to ATS-b.

Figure 6.5. Localization of ATS-b probe in P. falciparum infected red blood cells. Top panel: 100 nM Probe incubated for 30 min followed by 30 min of 1:200 dilution of avidin Alexafluor 647 fluorophore. Bottom panel: No probe control; avidin fluorophore only. Scale bar = 1 µm. Yellow arrows point to hemozoin.

The ATS-b probe appears to localize around the hemozoin (yellow arrows) which is within the DV of the parasite but appears to be both within the DV and the cytosol. Additional measurements can be done to determine exactly how much of the fluorescence is localized to the

DV versus the cytosol, and how this changes when exposing the iRBCS to competing ATS

(Figure 6.5). The probe fluorescence could also be measured in the presence of a secondary fluorophore to measure colocalization of the probe with that of a DV-specific fluorophore such as LysoSensor Green [Hartwig et al., 2009]. The Cooper group previously synthesized fluorescent artemisinin probes and measured cellular accumulation as well as colocalization with

125 several organelle specific fluorescent dyes, noting that their probes colocalized best with the vacuolar stain LysoSensor Green and BODIPY which stains neutral lipids (also frequently associated with areas containing hemozoin [Pisciotta et al., 2007]).

The subcellular localization of DHA-a was next analyzed similarly (see Section 2.2.10).

Briefly, cells were again fixed with formaldehyde and washed with PBS to remove non-adherent cells as with ATS-b, but DHA-a-treated cells were additionally coupled to azide-labeled Alexa

Fluor 488 before imaging.

Figure 6.6. Localization of DHA-a probe in P. falciparum infected red blood cells. Top panel: 100 nM alkyne probe incubated for 30 min followed by 30 min of 1:400 dilution of Azido-Alexa Fluor 488. Bottom panel: No probe control; Azido fluorophore only. Scale bar = 1 µm.

Figure 6.6 shows subcellular localization of the DHA-a probe. Importantly, incubation of trophozoites with Azido-Alexa Fluor 488 (but no alkynyl probe) yields no fluorescent signal within the iRBC (Fig 6.6, bottom panel).

Next, ATS-b was next tested vs. competition with underivatized ATS. While it is clear that the probe diffuses well into the parasite (Figure 6.5), for additional analysis of the probe it is

126 necessary to know if the probe targets are similar to that of underivatized ATS. If the two compounds bind the same targets, they will compete for binding space. Thus, if parasites are incubated with both biotinylated ATS and underivatized ATS, and since both the probe and ATS bind irreversibly, the fluorescence should decrease with increasing amounts of underivatized

ATS. Figure 6.7 shows parasite incubated with 100 nM probe and increasing molar equivalents of underivatized ATS. The top panel, incubated with 1:1 ATS-b : ATS, already shows dramatically reduced fluorescence showing that ATS competes very well vs. ATS-b and the biotinylated ATS probe therefore mimics the binding of the parent drug (ATS).

127

Figure 6.7. Competition experiments between biotinylated artesunate probe versus underivatized artesunate in P. falciparum infected red blood cells. (A) 100 nM Probe incubated for 30 min. (B) 100 nM probe co-incubated with 100 nM artesunate. (C) 100 nM probe followed by incubation with 100 nM artesunate. (D) 100 nM artesunate followed by incubation with 100 nM probe. (E) 1 µM artesunate followed by incubation with 100 nM probe. (F) 10 µM artesunate followed by incubation with 100 nM probe. All incubations of probe and drug followed by 30 min of 1:200 dilution of avidin fluorophore. Scale bar = 1 µm for all except (1), where scale bar is 2 µm. For each panel, all microscope settings were the same for collection of each image, thus each image can be directly compared.

128 Increasing amounts of competing ATS yield lower and lower levels of fluorescence within the parasite and also a more diffuse staining across the entire red blood cell. These data suggest that localization of ATS is pharmacologically relevant, and also that binding of the probe to intracellular targets is covalent as expected, as cells are washed extensively after incubation with

ATS-b (Chapter 2).

Lastly, localization of the DHA-a probe vs. time of incubation was inspected. As shown in Figure 6.8, the level of fluorescence within the iRBC is proportional to the time the parasites are exposed to 100 nM alkyne probe. This suggests that there are multiple parasite targets to which the probe can bind. Plateauing of the fluorescent signal within the iRBCs could suggest that the drug targets or spaces the probe could occupy become saturated over time.

Figure 6.8. Time-dependent localization of DHA-a probe. 100 nM alkyne probe incubated in infected red blood cells followed by 30 min of 1:400 dilution of Azido-Alexa Fluor 488. (A) 30 minute incubation (B) 15 minutes (C) 30 seconds (D) No probe control; azido fluorophore only. Scale bar = 1 µm.

129

6.3 Discussion

Biotinylated ATS (ATS-b) and alkynyl DHA (DHA-a) probes were synthesized in order to probe the subcellular localization of Art drug targets in P. falciparum parasites. Synthesis of these simple drug probes combined with high resolution spinning disk confocal microscopy

(SDCM) can better define drug localization, drug – target binding, and influx / efflux of these drugs. The clickable DHA-a synthesized in this study was similar in nature to probes synthesized earlier by the O’Neill group [Ismail et al., 2016a,b] and the Lin group [Wang et al., 2015], each of which include terminal alkynyl groups. However, to our knowledge, synthesis of ATS-b is novel and does not require additional chemical steps within the parasite in order to be visualized.

On the other hand, the longer tail introduces two potential problems, the first being that the drug might cross the parasite plasma membrane less easily, and the second being that the ATS tail could potentially be cleaved before interaction with the target and/or isolation on an avidin column. The alkynyl probe synthesized here might therefore ultimately prove to be more useful, though its clear that the ATS-b tail stayed in tact long enough to yield, bright and well defined fluorescence localization within the parasite.

Beyond describing syntheses of the probes, this study is novel for several reasons. First, previous work with Art drug probes [Ismail et al., 2016a; Wang et al., 2015] did not examine subcellular localization of the probes via microscopy. Second, previous research only examined probe targets for 3D7 parasites. 3D7 is a chloroquine-sensitive and Art-drug sensitive strain, whereas this study uses both CQS (HB3) and CQR (Dd2) P. falciparum strains. All ArtR strains developed from a chloroquine resistant (CQR) background, so CQR strains of P. falciparum may be more relevant for examining Art drug localization. Ideally, these experiments will be redone using now available ArtS and ArtR strains [Straimer et al., 2015]. At the time these experiments

130 were performed, such strains were not available. Targeted analysis of the P. falciparum proteome before and after drug probe exposure for ArtS and genetically mutated ArtR would ideally help define metabolic perturbations linked to drug sensitivity versus resistance.

Examining in more detail three previous Art proteomics studies [Ismail et al., 2016a,b;

Wang et al., 2015], Wang et al. found 124 potential protein targets to which their synthetic Art analog bound; Ismail et al. found 49 such targets, and 62 targets for their non-Art trioxane probe.

There are 19 proteins in common between the three studies (see Chapter 1). Of these, roughly

25% were vacuolar proteins, 15% were cytosolic, 10% were microtubule associated and the rest associated with different organelles or membrane-associated (Table 1.3). These results may suggest that Art drugs react somewhat randomly with cellular proteins. Art drugs, upon reacting with ferrous heme, immediately form oxygen centered radicals which convert to carbon centered radicals before covalently attaching to target proteins and other biologically relevant molecules

(Chapter 4). The lifetime of this conversion from oxy-centered radical to carbon-centered radical involving electron rearrangement is short, with a rate constant of > 1x107 s-1. [Butler et al.,

1998] If we consider the half life of a carbon centered radical as in Art to have a half-life on the microsecond scale [Ostdal et al., 1997; Neta et al., 2009], the radical could diffuse at least 500

µm in this time assuming an aqueous environment (this simplified model does not take into account the viscosity of membranes or organelles the radical may pass through). The diameter of a parasite within a red blood cell is roughly 4-6 µm (Figure 6.9), suggesting that an Art-drug carbon centered radical could reach any part of the parasite before decaying. Even if the half-life were one nanosecond, the adjusted distance would be 0.5 µm, so the radical would have time to travel about 1/10 of the way through the entire parasite, still being able to reach a substantial amount of proteins.

131 RPM PVM PPM RBC Cytosol

DV: ~ 1.75 µm Parasite: ~ 4 µm RBC: ~ 6 µm

Figure 6.9. Schematic representation of a trophozoite-infected red blood cell, drawn approximately to scale. Figure describes the red blood cell cytsol, red blood cell plasma membrane (RPM), parasitophorous vacuolar membrane (PVM), parasite plasma membrane (PPM), and parasite digestive vacuole (DV) drawn approximately to scale [Gligorijevic et al., 2006a; King 1988; Mehlhorn 1988].

With these concepts is mind, it is unclear if Art drug behavior would be any different for

ArtS vs. ArtR parasites unless conversion to Art drug radicals differed. Additional localization studies could be helpful to more fully test if Art drugs accumulate in certain subcellular compartments. Only about 25% of protein hits (see Chapter 1) in common between the three studies were vacuolar proteins. Co-localization studies with vacuolar stains as well as ER stains, nuclear stains, etc. could further highlight where these drug probes concentrate. Considering the likelihood that there are many possible targets for Art drugs to bind, it would make sense that Art drug binding to parasite targets increases with time, as seen in Figure 6.8.

The Art drug probes synthesized in this study have been shown to retain their cytostatic effectiveness as well as their ability to concentrate within parasites in the iRBC. They can now

132 be used for several additional key experiments. For example, using fluorescence microscopy and co-incubating parasites with both probe and markers for parasite compartments such as the DV, nucleus, etc., where the probe may specifically concentrate could be determined. Importantly, the probes could be used for detailed “omics” studies, comparing ArtS and ArtR parasites for a more complete picture of which Art-based drug adducts produced within the parasite could be the most pharmacologically relevant. This could include a full analysis of not only proteins, but also lipids and metabolites to which Art binds. This is particularly relevant as one target of Art is FPIX

(Chapter 4) and it is also known that Arts and similar trioxanes cause lipid peroxidation [Hartwig et al., 2009; Hartwig et al., 2011]. Comparing the Art drug binding targets for ArtS vs. ArtR parasites could allow a grouping of targets by category and could highlight types of targets that might be most associated with the ArtR phenotype. The affinity purification processes would be different for proteins vs. metabolites and lipids, where the latter two categories could be extracted using a 1:3:1 mix of CHCl3:MeOH:H2O where lipids should be extracted into the organic phase and metabolites in the aqueous phase [Olszewski et al., 2009; Cobbold et al.,

2016; Allman et al., 2016]. Taking into account the findings from previous proteomics studies

(Chapter 1) [Ismail et al., 2016a,b; Wang et al., 2015] we might predict a significant amount of vacuolar metabolites and lipids to bind to these Art drug probes. It would also be interesting to see if FPIX is found to bind the Art probes, and if there is any significant difference in Art binding for ArtS vs. ArtR parasites as seen previously (Chapter 4) [Heller et al., 2018b].

133 CHAPTER 7. CONCLUSIONS

The most current records from the WHO list an estimated 219 million clinical cases and an estimated 0.5 million deaths occurring annually due to malaria [WHO 2017]. Despite the advent of new antimalarials, drug resistance has been reported for virtually every known antimalarial to varying degrees. The initial success of ACTs as frontline antimalarials showed great promise, however ACT resistance is emerging in some regions, and is typically defined clinically as a “delayed clearance phenotype” (DCP). Because of the declining efficacy of current ACTs and the need for new ACT partner drugs, the exact mechanism for activation of

Art based drugs and the relation to parasite death remain an active area of research. Definition of

ACT molecular targets in P. falciparum is critical for the development of improved ACTs that contain either a modified endoperoxide drug, different partner drug(s), or both.

As described in detail in Chapter 1, many antimalarials target the hemoglobin digestion pathway. This process involves sequential plasmepsin and falcipain degradation of hemoglobin

(Hb) to free amino acids and FPIX heme. The parasite can utilize amino acids but must remove or detoxify ferric FPIX which is toxic to the parasite. In order to avoid this fate, the parasite sequesters free FPIX into an inert crstyalline head-to-tail dimer called hemozoin (Hz).

Antimalarial drugs such as the quinolines act by inhibiting this crystallization process and causing free toxic FPIX to build up in the parasite digestive vacuole (DV). The Roepe lab has described the interaction between free FPIX and quinolines in great detail [Leed et al., 2002; de

Dios et al., 2003; Gorka et al., 2013c and others]. Formation of a dative complex between drug and heme can prevent the crystallization process [Marques et al., 1996; de Villiers et al., 2012;

Olafson et al., 2015]. Non-covalent binding of quinolines to dimeric FPIX is also strong enough

134 to inhibit the self-association of FPIX, providing a second, simple yet efficient way to inhibit growth and fluctuation of the parasite [Leed et al., 2002; Gorka et al., 2013c].

With this in mind, I set out to study the interactions between FPIX and Art-based antimalarials. Art, derived from the Chinese herb Artemisia annua, contains a highly reactive

1,2,4 trioxane ring. In its reduced form, free FPIX is known to catalyze endoperoxide cleavage of

Art-based drugs. As described in Chapters 1 and 4, the activated drug likely proceeds from an oxy radical form to a carbon centered radical form that is then capable of alkylating a variety of drug targets within malarial parasites. It was previously established by the Meshnick, Posner, and

Meunier groups that Arts bind reduced FPIX covalently and irreversibly in vitro (Chapters 1 and

4). It has also been proposed for some time that FPIX liberated upon Hemoglobin catabolism is a target of Arts within the parasite [Robert et al., 2005; Creek et al., 2008; Zhang et al., 2008;

Zhang et al., 2009; Mercer et al., 2011; Wang et al., 2015]. The most biologically relevant means of activating DHA and other ART-based drug endoperoxide groups to create carbon- centered radicals capable of alkylating Art targets is via Fe(II)-catalyzed Fenton chemistry.

[Posner et al., 1992; Robert et al., 2001]. In the malarial parasite, the most abundant source of

Fe(II) is Fe(II)PPIX, released upon Hemoglobin catabolism. Free Fe(II)PPIX does not exist in unifected RBCs, as breakdown of Hemoglobin by the parasite or some source is required. Free

Fe(II)PPIX activates the Art endoperoxide as effectively as does Free Fe(II) [Hassett et al.,

2018b]. Additionally, levels of FPIX reach up to 4-5 mM within the malarial parasite [Heller et al., 2018a]. Thus, I suggest that free FPIX is the primary activator of Art drugs within malarial parasites. If this is the case, we might expect to find differences between the levels of FPIX or

FPIX-Art adduct within ArtS vs. ArtR parasites.

135 In Chapter 3, I used optimized extraction procedures and mass spectrometry to quantify the abundance of FPIX and hemozoin at various stages of the P. falciparum life cycle. Intially,

Kyle et al., 2015 proposed that the ring stage of DCP parasites was extended relative to the control. In this study, growth of DCP strains was compared to that of a non-isogenic laboratory control CQR strain isolated much earlier (W2). I sought to expand on this with newly available isogenic parasites expressing WT vs. DCP-associated K13 isoforms [Straimer et al., 2015].

Indeed, I found that K13 mutations were associated with an extended ring stage and a truncated trophozoite stage. This led us to pose the question, if the trophozoite stage is truncated in the

ArtR parasite, is the parasite unable to digest the same amount of Hemoglobin? And if so, would that lead to less available target for Art drugs? To answer these questions, I devised an assay to extract purified heme from bulk parasite culture. After perfecting this assay, I isolated free FPIX and hemozoin at various stages of the P. falciparum life cycle in order to compare levels of both forms of FPIX for ArtR vs. ArtS parasites. Abundance of free FPIX vs. time for ArtS parasites was as expected, with low levels of FPIX detected in the ring stage and a distinct peak corresponding to Hemoglobin digestion in the mid – late trophozoite stage. For ArtR parasites, however, the phenotype was very different, with a higher baseline for FPIX abundance across the entire life cycle and a much broader, shallower peak at the trophozoite stage. Interestingly, maximal FPIX abundance peaked at the same time for ArtR and ArtS parasites. Maximal levels of hemozoin were similar for ArtR and ArtS parasites as well, but hemozoin formation began sooner for ArtR, which makes sense considering the higher levels of ring stage free FPIX for

ArtR parasites. This phenotype for ArtR vs. ArtS levels of free FPIX and hemozoin held true for two additional strains with WT and DCP-associated K13 mutations. In sum, these results suggest

136 that the levels of free FPIX and hemozoin, and likely the Hemoglobin catalytic process, differ for

DCP parasites commensurate with their altered red blood cell life cycle.

This story is continued in Chapter 4, where the heme extraction protocol was adapted for isolation of FPIX-Art adduct from the parasite. If there is less free FPIX available for ArtR parasites, this suggested that there is less target for Art drugs to bind, thus leaving the drug less potent. We sought to extract and quantify FPIX-DHA adduct from DHA-dosed parasites in order to compare formation of adduct in ArtS vs. ArtR parasites. For model studies, in vitro adduct was made by incubating FPIX, GSH, and DHA in 40% DMSO for three hours and exposing the adduct to SDS and guanidinium chloride to mimic the FPIX extraction process used for live parasites. MS data for variable amounts of in vitro adduct was then used to calibrate concentration of adduct vs. peak area. Once a calibration curve could be designed, parasites were bolus dosed with DHA for 6 hr and FPIX-DHA adducts were extracted and quantified by LC-

MS. As might have been expected, there were significantly lower levels of FPIX-DHA adduct for ArtR parasites as compared to ArtS, at the trophozoite stage of parasite development because less free FPIX is found for ArtR at this stage. However, surprisingly, levels of FPIX-DHA adduct were lower for ArtR ring and trophozoite stages as well. While the relationship between decreased FPIX-DHA adduct across all stages in ArtR with ring-stage sensitivity (as quantified by the RSA [Witkowski et al., 2013; Straimer et al., 2015]) is not immediately clear from these data, I propose that the FPIX-DHA adduct could affect metabolic processes specific to the ring stage. These might include ring stage membrane biogenesis or membrane development, upregulated transcription and translation that occurs at the ring stage, or early ring stage

Hemoglobin catabolism that may differ from catabolism in the trophozoite stage. Additionally, while we see a difference between both the abundances of free FPIX and the amounts of FPIX-

137 DHA adduct formed within the parasite for ArtS vs. ArtR parasites, the adduct abundance corresponds to only around 1% of the total amount of free FPIX present in the parasite. This supports the hypothesis that activated Art promiscuously targets many small molecules and proteins, as described in Chapter 6.

Chapter 5 further explores the interaction between an Art-based drug (ATS) and

Fe(II)PPIX in vitro via NMR spectroscopy and additional mass spectrometry experiments.

Specifically, we monitor the reaction between Fe(II)PPIX and ATS +/- various quinoline based drugs which are also known to bind to FPIX. In these experiments, Fe(II)PPIX is generated from

Fe(III)PPIX via reduction by 5X molar excess glutathione. The rate of ATS activation and glutathione oxidation is then observed by NMR upon addition of ATS. As described in Chapter

1, while the mechanism of action of Art drugs remains controversial, it is thought that FPIX is involved in the activation of the endoperoxide group, which can cause a cascade of radical products to alkylate protein, lipid, and small molecule targets within the parasite. In view of this, the chemical reactions studied in Chapter 6 are important for better understanding this important class of drugs. In this study, the rate of reaction between FPIX and ATS is specifically monitored via the rate of loss of the 3-CH3 proton signal for the methyl group directly adjacent to the endoperoxide bridge. Upon schism of the endoperoxide, the methyl group becomes part of an acetyl group which leaves the molecule as free acetate. Thus, loss of the 3-CH3 proton directly corresponds to breakage of the endoperoxide and the overall reaction of ATS with FPIX. We find that conversion of ATS to the FPIX-ATS complex is indirectly proportional to oxidation of

GSH into GSSG. The amount of unreacted ATS (i.e. size of 3-CH3 proton signal) was monitored as a function of time in the absence vs. presence of other FPIX – binding quinoline antimalarial drugs, and quinolines were found to interact preferentially with the FPIX, slowing the reaction

138 with ATS. Reaction with ATS was impeded with all quinolines except for 9-epi-quinine, which binds much more weakly to FPIX and in a different manner, i.e. in edge - to - face fashion

[Gorka et al., 2013c]. The presence of epi-quinine actually promoted FPIX to form a 2:1

FPIX:ATS complex. That conversion of ATS to the FPIX-ATS complex is slowed in the presence of additional traditionally FPIX – binding antimalarials suggests that the activation of

ATS is dependent on access to free ferrous iron within Fe(II)PPIX.

While the in vitro evidence suggests Art drugs are activated by (and bind covalently to)

FPIX, recent proteomics studies suggest that there could also be several dozen protein targets for

Art drugs, though across the three studies there are only 19 overlapping targets [Wang et al.,

2015; Ismail et al., 2016a,b]. This suggests that alkylation of targets could be somewhat random.

However, to our knowledge there has not been a thorough comparison of protein targets for ArtS vs. ArtR parasites. While these additional potential targets of Art drugs are interesting, what might be of greater interest is how DCP affects the targets and the abundance of these additional drug-target complexes. With that information we could create a better picture of what targets might be more important or relevant to the mechanism of Art drug resistance.

In Chapter 6, I synthesized two Art derivatives, one a biotinylated ATS probe and one a clickable alkynyl Art which could then be coupled to an azido fluorophore as well as a biotin azide. Both probes’ subcellular localization was visualized using fluorescence confocal microscopy, with the end goal in the future being potential proteomics or lipidomics studies using one or both probes. Both probes were tested for their effectiveness against P. falciparum in

IC50 assays compared with respective underivatized drugs to which they were most similar, i.e.

ATS and ATM. Each probe retained its biological activity, as IC50 values vs. HB3 and Dd2 were comparable to that of underivatized Arts. After determining that the probes had retained

139 biological activity, the subcellular localization of each probe was analyzed using spinning disk confocal microscopy. Each probe localized to the iRBC and, in each case, incubation of cells with fluorophore but no probe did not lead to fluorescence localization inside the parasite or red blood cell, suggesting that binding to probe is necessary for each fluorophore labelling. In order to determine if the probes’ targets are similar to that of underivatized drugs, ATS-b was tested vs. competition with underivatized ATS. Since both the probe and ATS bind irreversibly, competition for binding sites should theoretically lead to a loss of signal. Indeed, fluorescence from the ATS-b probe decreased when probe was co-incuabted with increasing amounts of underivatized ATS. Lastly, localization of the DHA-a probe vs. time of incubation was inspected, where the DHA-a was seen to concentrate within the iRBC in a time-dependent manner, in agreement with the hypothesis that Art drugs bind to many targets within the parasite.

The methods described in this thesis can be used to easily quantify levels of FPIX and

FPIX-drug adducts in an unprecedented manner. The importance of our finding a way to isolate and quantify not only free FPIX but also a FPIX-Art adduct from within the parasite must be stressed. Previously, parasite FPIX has only been quantified via UV-Vis, often relying on indirect measurements such as FPIX binding to pyridine and quantifying pyridine absorbance

[Ncokazi et al., 2005]. Measuring the unique kinetics for the digestion of Hemoglobin into free

FPIX in ArtR vs. ArtS parasites, and in turn levels of FPIX-DHA adduct, is entirely novel. Many previous studies have offered support for the hypothesis that Art drugs’ mechanism of action is reliant on endoperoxide activation via Fe(II) of some form; however, most of these studies have focused on either in vitro work [Posner et al., 1992; Hong et al., 1994; Meunier et al., 2002a] or, more indirect measurements such as adding inhibitors of Hemoglobin digestion and seeing decreased Art potency in parasites [Klonis et al., 2011]. In sum, this work aims to provide a

140 more complete picture of Art drug-target interactions with FPIX and other potential protein targets, as well as how that relates to the ArtR. By quantifying abundance of free FPIX and

FPIX-DHA adducts, monitoring Art-FPIX interactions via UV-Visible and NMR spectroscopies, and synthesis of Art derivatives for visualization of subcellular localization using fluorescence confocal microscopy as well as future proteomics studies, the work presented in this thesis has illuminated unknown aspects of Art pharmacology as it pertains to FPIX interactions and DCP.

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