Towards functional assignment of
Plasmodium membrane transport proteins:
an experimental genetics study on four diverse proteins
A thesis submitted for the degree of Doctor of Philosophy of The Australian National University and Humboldt University of Berlin
by François Laurent Frederic Korbmacher
Submitted July 2020
© Copyright by François Laurent Frederic Korbmacher, 2020 All Rights Reserved
Declaration
The research presented in this thesis, except where otherwise acknowledged, is my own original work. Research conducted in cooperation is noted.
The study was performed at the Australian National University under supervision of Prof. Dr. Alex G. Maier and at the Humboldt University of Berlin under supervision of Prof. Dr. Kai Matuschewski within the frame of a Cotutelle-degree by the IRTG2290 graduate program.
No portion of the work presented in this thesis has been submitted for another degree.
The word count for this thesis excluding tables, figures, bibliographies and appendices is 43,156 words.
François Laurent Frederic Korbmacher
Berlin, July 2020
Towards functional assignment of
Plasmodium membrane transport proteins:
an experimental genetics study on four diverse proteins
Dissertation
Zur Erlangung des akademischen Grades Doctor of Philosophy (PhD) im Fach Biologie
eingereicht an der Lebenswissenschaftlichen Fakultät der Humboldt- Universität zu Berlin und der Australian National University
von François Laurent Frederic Korbmacher geb. am 20.06.1990 in Neustadt an der Weinstraße
Präsidentin der Humboldt-Universität zu Berlin Prof. Dr.-Ing. Dr. Sabine Kunst
Dekan der Lebenswissenschaftlichen Fakultät Prof. Dr. Bernhard Grimm
Gutachter: Vorsitz: Prof. Dr. Christian Schmitz-Linneweber 1. Prof. Dr. Kai Matuschewski 2. Prof. Dr. Alex Maier 3. Prof. Tania De Koning-Ward 4. Prof. Dr. Simone Reber 5. Dr. Jandeli Niemand Tag der mündlichen Prüfung: 10.12.2020
Summary
Membrane transport proteins (MTPs) transfer nutrients, metabolic products and inorganic ions across membranes. Many MTPs are essential for Plasmodium blood infection and gain importance as candidate drug targets in malaria therapy, whereas the physiological functions of many MTPs still remain enigmatic. In this thesis, we applied experimental genetics to determine key characteristics of four selected Plasmodium MTPs, including spatio-temporal expression and phenotypical analysis of knockout mutants. We employed the murine malaria model parasite Plasmodium berghei and in vitro blood cultures of the human malaria parasite Plasmodium falciparum. For this study, we selected one conserved MTP called FT2, which was previously shown to transport folate, a P-type ATPase that is specific for P. falciparum as well as two essential MTPs, CRT and ATP4, with important roles in anti-malarial therapy. These targets exemplify the range of druggable candidates and illustrate the potential and limitations of reverse genetics to decipher their physiological roles for Plasmodium life cycle progression.
A combination of transgenic and knockout strategies was applied to the P. berghei folate transporter 2 (FT2). We show that endogenously tagged FT2 localises to the apicoplast membranes, and is broadly expressed throughout the parasite’s life cycle. Strikingly, in two stages, schizonts and sporozoites, FT2 displays a dual localisation. Analysis of FT2-deficient parasites revealed a severe sporulation defect in the vector; the vast majority of ft2– oocysts form large intracellular vesicles which displace the cytoplasm. Accordingly, very few sporozoites are generated and these are non-infectious to the mammalian host, resulting in a complete arrest of Plasmodium transmission. A candidate aminophospholipid P-type ATPase, that is encoded by human malaria parasites, but not P. berghei and related Vinckeia parasites, was assessed by a CRISPR/Cas9-mediated gene disruption. Compared to many vital P-type ATPases this gene is dispensable for asexual blood replication. Two MTPs, ATP4 and CRT are prime targets for antimalarial therapies. A comprehensive spatio-temporal expression analysis of transgenic parasites expressing mCherry- tagged proteins revealed expression beyond blood infection, indicative of functions in additional parasite stages.
Together, the findings of this study contribute towards a better understanding of the roles of the four MTPs based on localisation, expression and functional deletion.
i
Zusammenfassung
Membran Transport Proteine (MTPs) transferieren Nährstoffe, metabolische Produkte und anorganische Ionen über Membranen. Da etliche MTPs in den Plasmodium Blutstadien essentiell sind, geraten sie zunehmend in den Fokus der Wirkstoffentwicklung. Die eigentlichen physiologischen Rollen der Transporter sind jedoch oft ungeklärt. In dieser Arbeit wurden mittels experimenteller Genetik funktionelle Charakteristika, wie zeitliche und räumliche Expressionsmuster sowie phänotypische Analysen von Mutanten untersucht. Am Maus Parasiten Plasmodium berghei und der Plasmodium falciparum Blutstadien-Kultur wurden vier MTPs ausgewählt: ein konservierter Folat Transporter (FT2), sowie eine P. falciparum- spezifisches P-Typ ATPase und zwei essentielle MTPs mit wichtigen Rollen in der Malaria-Therapie (CRT und ATP4). Diese Auswahl verkörpert ein breites Spektrum an MTP Kandidaten für Wirkstoffziele und reflektieren zudem das Potenzial und die Grenzen funktioneller Analysen von Plasmodium MTPs mittels reverser Genetik.
Für den Folat Transporter 2 (FT2) wurde eine Kombination von transgenen Strategien auf P. berghei angewandt. Durch ein endogenes tag von FT2 wurde die Lokalisierung im Apicoplast, eine doppelte Lokalisierung in Schizonten und Sporozoiten, sowie dessen Expression über fast den kompletten Zyklus hinweg gezeigt. Nach der Deletion von FT2, wiesen die Parasiten einen Defekt während der Sporulation im Vektor auf: der Großteil der ft2– Oozyten formt große, intrazelluläre und Zytoplasma verdrängende Vesikel. Demzufolge bilden sich nur sehr wenige und nicht infektiöse Sporozoiten, was letztendlich zur Unterbrechung des Lebenszyklus der Parasiten führt. Eine vermeintliche Aminophospholipid P-Typ ATPase, für die kein kodierendes Gen in der P. berghei mit inbegriffenen Vinckeia Klade auftritt, wurde mittels CRISPR/Cas9 in P. falciparum genetisch deletiert und die Mutante analysiert. Im Gegensatz zu den meisten vitalen P-Typ ATPasen erweist sich das Gen in den asexuellen Blutstadien als entbehrlich. Des Weiteren bilden die MTPs ATP4 und CRT einen einflussreichen Faktor bei Malaria-Therapien. Eine umfassende Analyse von räumlichen und zeitlichen Expressionsmustern von transgenen Parasiten mit mCherry-getaggten Proteinen zeigt ein Expression der beiden MTPs über die Blutstadien hinaus, was auf zusätzliche Funktionen in den jeweiligen Stadien verweist.
Diese Studie trägt, basierend auf Lokalisation, Expression und funktioneller Deletion, zur funktionellen Entschlüsselung der vier untersuchten MTPs bei.
ii Table of contents
1 st Chapter: General Introduction ...... 1
1.1 Plasmodium and malaria ...... 1
1.2 Life cycle of the Plasmodium parasite ...... 1
1.3 Membrane Transport Proteins in Plasmodium ...... 4
1.4 Towards a functional classification of MTPs ...... 6
1.5 Aims of the study ...... 8
2 nd Chapter: Materials and Methods ...... 9
2.1 Materials ...... 9 2.1.1 Experimental organisms ...... 9 2.1.2 Plasmodium targeting vectors ...... 9 2.1.3 Enzymes ...... 10 2.1.4 Antibodies ...... 10 2.1.5 Media, buffers, stock and working solutions...... 10 2.1.6 Laboratory equipment ...... 11 2.1.7 Consumables ...... 12 2.1.8 Commercial kits ...... 13 2.1.9 Chemicals ...... 13 2.1.10 Software and databases ...... 15 2.2 Methods ...... 15 2.2.1 Molecular- and microbiological methods for P. berghei and P. falciparum ...... 15 2.2.1.1 Polymerase Chain Reaction (PCR)...... 15 2.2.1.2 Agarose gel electrophoresis ...... 16 2.2.1.3 Digest of DNA with restriction enzymes ...... 16 2.2.1.4 Ligation of plasmids and DNA fragments ...... 17 2.2.1.5 Preparation of electrocompetent PMC103 E. coli ...... 17 2.2.1.6 Transformation in competent E. coli ...... 18 2.2.1.7 Plasmid DNA isolation ...... 18 2.2.1.8 Molecular cloning with Gibson assembly ...... 19 2.2.1.9 Isolation of RNA with Trizol and Glycogen ...... 19 2.2.1.10 Sanger Sequencing ...... 20 2.2.1.11 Isolation of genomic DNA ...... 20 2.2.1.12 DNA precipitation ...... 20 2.2.1.13 Complementary DNA (cDNA) conversion by reverse transcription...... 20 2.2.1.14 Quantitative real-time PCR (qPCR) ...... 20 2.2.2 Live fluorescence microscopy ...... 21 2.2.3 Transfections of recombinant P. berghei lines and phenotyping ...... 22
iii 2.2.3.1 Generation of recombinant P. berghei strain ANKA parasites lines ...... 22 2.2.3.2 Transfection of P. berghei schizonts ...... 24 2.2.3.3 P. berghei schizont culture ...... 25 2.2.3.4 Selection of isogenic P. berghei cell line ...... 25 2.2.3.5 Cryopreservation of mouse blood infected with P. berghei...... 26 2.2.3.6 Isolation of P. berghei from infected mouse blood ...... 26 2.2.4 P. berghei life cycle analysis ...... 26 2.2.4.1 Giemsa-stain of thin blood smears ...... 26 2.2.4.2 Asexual blood stage development...... 27 2.2.4.3 Sexual blood stage development ...... 27 2.2.4.4 Ookinete formation culture ...... 28 2.2.4.5 Breeding of Anopheles stephensi mosquitoes ...... 28 2.2.4.6 Transmission of blood stage parasites to Anopheles stephensi mosquitoes ...... 28 2.2.4.7 Analysis of parasite-infected mosquito midguts ...... 29 2.2.4.8 Sporozoite extraction of mosquitoes’ salivary glands ...... 30 2.2.4.9 Transmission of sporozoites to naïve mice ...... 30 2.2.4.10 In vitro infection of hepatoma cells (Huh7) with sporozoites ...... 30 2.2.5 Transfection of recombinant P. falciparum 3D7 lines ...... 31 2.2.5.1 Generation of recombinant P. falciparum 3D7 lines ...... 31 2.2.5.2 Maxi preparation of DNA ...... 32 2.2.5.3 Preparation of plasmid for transfection...... 33 2.2.5.4 Transfection of P. falciparum rings stages ...... 33 2.2.5.5 Diagnostic PCR for screening of recombinant P. falciparum ...... 36 2.2.5.6 Isolation of clonal parasite line based on limiting dilution...... 36 2.2.6 In vitro culturing of asexual P. falciparum 3D7 stages ...... 37 2.2.6.1 Preparation of uninfected erythrocytes...... 37 2.2.6.2 Serum deactivation ...... 37 2.2.6.3 In vitro culture of P. falciparum ...... 37 2.2.6.4 Assessment of parasitemia in an in vitro culture of P. falciparum ...... 38 2.2.6.5 Thawing of P. falciparum stabilates ...... 38 2.2.6.6 Saponin lysis ...... 39 2.2.6.7 Cryopreservation of P. falciparum parasites lines ...... 39 2.2.7 P. falciparum SYBR safe-based in vitro growth assay ...... 40 3 rd Chapter: An apicoplast-resident folate transporter is essential for sporogony of malaria parasites ...... 41
3.1 Introduction ...... 41
3.2 Results ...... 43 3.2.1 FT2 is broadly expressed throughout the life cycle and localises to the apicoplast ...... 43 3.2.2 Deletion of FT2: normal blood propagation but arrest of transmission by loss of sporulation. ....46
iv 3.2.3 Rescue of defect by ectopic expression of FT2::mCherry in SIL6 in FT2-deficient and wild type parasites...... 52 3.2.4 Heterozygous oocysts after cross-fertilisation of ft2– and wild type parasites overcome defect and produce haploid ft2– sporozoites ...... 56 3.2.5 FT2 ortholog in P. falciparum ...... 59 3.2.6 Methionyl-tRNA formyltransferase (FMT): a folate dependent pathway in the apicoplast ...... 62 3.3 Discussion ...... 67
4 th Chapter: Role of a putative aminophospholipid ATPase (Pf3D7_1468600) with an ortholog loss in rodent malaria parasites ...... 71
4.1 Introduction ...... 71
4.2 Results ...... 72 4.2.1 Phylogenetic analysis of Pf3D7_1468600...... 72 4.2.2 CRISPR/Cas9-mediated disruption of Pf3D7_1468600 ...... 72 4.2.3 SYBR-safe growth assay indicates that Pf3D7_1468600 is involved in growth-efficiency in the asexual blood stages ...... 75 4.3 Discussion ...... 76
5 th Chapter: Expression profiling of two vital P. berghei membrane transport proteins, CRT and ATP4 ...... 79
5.1 The Chloroquine Resistance Transporter ...... 79 5.1.1 Introduction ...... 79 5.1.2 Results CRT ...... 81 5.1.2.1 An isogenic CRT::mCherry cell line was used for spatio-temporal analysis...... 81 5.1.2.2 Food vacuole resident CRT::mCherry co-localises with hemozoin crystals...... 82 5.1.2.3 CRT::mCherry shows an expression pattern beyond blood stages...... 85 5.1.2.4 Validation of CRT::mCherry signal quantification by stage-specific mRNA expression profiling ...... 87 5.1.3 Discussion CRT ...... 87 5.2 ATP4...... 90 5.2.1 Introduction ...... 90 5.2.2 Results ATP4 ...... 92 5.2.2.1 An isogenic ATP4::mCherry cell line was used for spatio-temporal analysis...... 92 5.2.2.2 ATP4 localises in parasite plasma membrane throughout life cycle progression with a new target in midgut sporozoites ...... 95 5.2.2.3 ATP4 shows expression throughout entire life cycle progression ...... 95 5.2.2.4 ATP4 mRNA expression ...... 96 5.2.3 Discussion ATP4 ...... 97 5.3 Technical discussion ...... 99
6 th Chapter: General Discussion ...... 102
v 6.1 Findings of this study ...... 102
6.2 Experimental genetics in Plasmodium parasites ...... 103
6.3 The transporter dilemma ...... 104
7 th Chapter: Conclusion and Outlook ...... 107
Literature...... 108
Acknowledgements...... 123
Abbreviations ...... 126
Appendix ...... 130
vi vii viii Towards functional assignment of
Plasmodium membrane transport proteins: an experimental genetics study on four diverse proteins 1ST CHAPTER - GENERAL INTRODUCTION
1st Chapter: General Introduction 1.1 Plasmodium and malaria
Plasmodium spp. are obligate intracellular parasites and belong to the order Haemosporida, which consists of over 500 parasitic species from at least 15 genera. All of them infect vertebrates, and are transmitted by blood-feeding dipteran vectors [1]. Haemosporidia are a member of the large phylum Apicomplexa, the only taxonomic group whose members are entirely parasitic and carry a four-layer membrane-bounded plastid: the apicoplast [2]. In the recent two decades, it has been recognised that Apicomplexans had a photosynthetic ancestry but lost photosynthesis during phylogenesis [3, 4]. The apicoplast is one of the characteristic features of the Apicomplexan phylum (besides Cryptosporidium), which parasitises the animal kingdom for more than 500 million years [5, 6].
Plasmodium spp. infect a broad range of vertebrate hosts, from reptiles to primates. Five species are known to infect humans on whose genome they applied high selection pressure since their early interactions. The most relevant example is Plasmodium falciparum, the etiological agent of malaria tropica, which causes the most virulent and prevalent form of human malaria [7, 8].
The virulence of a P. falciparum infection is mainly driven by the parasite’s continuous multiplication inside the erythrocytes, which causes the major clinical symptoms like fever-chill periodicity, fatigue, nausea and headache. The morbidity and mortality are associated with a broad range of pathologies, such as severe anaemia, multiorgan failure or coma. Correlating to the presence of the parasite and its Anopheles vector, the disease claims around half a million deaths every year [7].
1.2 Life cycle of the Plasmodium parasite
During the blood meal of the Anopheles vector, sporozoites are injected into the dermis of the human host. In transmission models it was shown that up to 1,000 sporozoites can be experimentally transmitted during extended blood meals [9]. However, only around 10-20% of the sporozoites enter the blood stream, which constitutes the first bottle neck the parasite encounters [10]. After sporozoites penetrate a blood vessel to reach the peripheral circulation, they travel through the blood stream until they reach the liver (Fig. 1 A). They cross the liver sinusoidal barrier
1 1ST CHAPTER - GENERAL INTRODUCTION
A STAGES LIVER M EROSOME
SALIVARY GLANDS
LIVER
POROZOITES E S MEROZOITE
M
AMMALIAN
POROZOITES M SCHIZONT S OSQUITO
RING STAGE H B
OST V TROPHOZOITE
ECTOR D OOCYSTS BLOOD STAGES
OOKINETE GAMETOCYTE C
EXFLAGELLATION MIDGUT Fig. 1 Plasmodium life cycle. Red arrows: transmission between vector and host. A: sporozoites infect liver cells and develop into liver stages and the merosome. Merosome escapes liver cells, ruptures and releases merozoites to blood circulation. B: merozoites invade erythrocytes and develop in asexual cycle to ring stages, trophozoites, schizonts and segmented merozoites which reinvade erythrocytes. C: a proportion of trophozoites commits to male (grey) and female (blue) gametocytes (shape represents P. falciparum cellular morphologies). After an uptake to the mosquito midgut, gametocytes get activated. Male gametocytes exflagellate and form microgametes which recombine with macrogametes. A new formed ookinete penetrates the mosquito midgut wall and colonises the midgut by embedding in the basal lamina. D: oocysts develop and form sporozoites. Mature sporozoites escape from oocyst and migrate to the mosquito’s salivary glands. With a new blood meal (E), sporozoites get injected in new host where they migrate to blood vessels and reach the liver through the circulation.
2 1ST CHAPTER - GENERAL INTRODUCTION and reach hepatocytes after traversing multiple cells. With this, the parasite escapes the clearance by Kupffer cells and forms a transient vacuole [11, 12]. Triggered by liver-cell factors, the sporozoite stops migration and invades a final liver cell to develop into a so-called exo-erythrocytic form (EEF) [13]. Two – ten days after the invasion, the EEFs perform intense structural replications and finally release a vesicle named merosome in the blood stream, filled with several thousands of merozoites [14]. Released merozoites circulate through the blood stream, and attach to erythrocytes through the formation of a tight junction between parasite and host-cell surface proteins [15] (Fig. 1, B). By this irreversible binding, the merozoite actively pushes itself in the erythrocyte, through forces generated by an actomyosin motor [16]. After the erythrocytic infection is established, the parasite forms ring stages, develops into trophozoites and finally to schizonts which contain around 30 daughter merozoites. The asexual cycle takes approx. 48 hours in P. falciparum and 24 hours in the rodent parasite Plasmodium berghei [17, 18]. A proportion of the asexual parasites commits to sexual development, driven by a distinct pattern of transcription factors and epigenetic regulators, which results in the formation of male and female gametocytes [18] (Fig. 1 C). Furthermore, it is discussed that environmental parameters trigger the induction of gametocytogenesis [19]. Gametocytes show developmental differences between P. berghei and P. falciparum. This is reflected by morphology, maturation time and location in the host [18]. Despite the difference, in both species gametocytogenesis is essential for the transmission to the mosquito, since gametocytes recombine in the mosquito’s midgut. Following activation by mosquito midgut parameters, like temperature or exposure to midgut acids, male and female gametocytes activate and egress from the host cell. The female gametocyte forms to a macrogamete, while the male gametocytes undergo exflagellation and form eight motile microgametes. Microgametes fertilise macrogametes, and the resulting zygotes transform into infective and motile ookinetes [20]. By active penetration of the mosquito’s midgut wall, the ookinetes colonise the midgut by embedding in the basal lamina of the midgut [21] (Fig. 1 D). Oocysts are formed and grow into large encapsulated spheres with a diameter of ~50 µm. They undergo several rounds of nuclear division followed by a concerted and tightly synchronized cytokinesis programme which results in the formation of thousands of sporozoites [22, 23]. The process of sporulation begins with the detachment of the plasma membrane from the oocyst capsule [24]. By continued vesicle fusion, the plasma membrane forms large
3 1ST CHAPTER - GENERAL INTRODUCTION fissures to ultimately yield the sporoblasts, which are isolated islands of parasite cytoplasm. Here, parasite organelles align below the plasma membrane from where the sporozoites emerge through a budding mechanism [25]. Upon protease-mediated rupture of the oocyst capsule [26], these slender and highly motile parasite stages migrate to the salivary glands from where they are transmitted during the mosquito’s blood meal (Fig. 1 E).
The vector of Plasmodium spp. are mosquitoes of the genus Anopheles. While male and female mosquitoes feed on sugar-rich liquids, only the female imago is seeking a blood meal to induce oviposition. Eggs develop in an aquatic phase via four larval stages to pupae, which hedge again to the adult imago [27].
1.3 Membrane Transport Proteins in Plasmodium
The Plasmodium parasite incorporates multiple membrane systems through membrane-bound compartments which are enclosed by up to four lipid-bilayers, as for instance in the Apicomplexa-specific apicoplast. Dependent on the environment during the parasite’s life cycle progression, Plasmodium harbors particular membranous structures like the digestive vacuole and the parasitophorous vacuole, which is closely linked to the parasite’s plasma membrane in intracellular stages [28]. During the intracellular phase, the parasite is also surrounded by the host cell’s plasma membrane. Such a multilayered ‘life style’ requires an appropriate trans-membranous transport machinery of substrates, which is mediated by Membrane Transport Proteins (MTPs) [29].
MTPs come in different types of proteins or protein complexes which form channels, carriers and pumps (Fig. 2 a). Channels form aqueous pathways, through which cargos of specific size and charge travel across the membrane at rapid rates. In contrast, carriers bind the substrate and transport the cargo driven by conformational changes across the bilayer. This can occur down an electrochemical gradient of the cargo (uniport), or against the electrochemical gradient driven by the gradient of another cargo which is either co-transported (symport) or transported in the opposite direction (antiport). To allow a gradient-independent transport, energy utilising pumps can transport substrates against the gradient by the hydrolysis of ATP. The active transport occurs at slower transport rates compared to channels [30]. Depending on
4 1ST CHAPTER - GENERAL INTRODUCTION
(a) (b) functional data putative function + + - unknown function gradient
ADP + P ATP - i - + uniporter anitiporter symporter 144 MTP genes Channels Pumps Carriers Fig. 2 Membrane Transport Proteins (MTPs) in Plasmodium. (a) Scheme of the different MTP types: channels (yellow), pumps (green) and carriers (blue). Gradient indicated by plus (+) and minus (-). Channels transport substrate down the gradient. Pumps can actively transport a cargo against the gradient by the hydrolyses of ATP. Carriers undergo a conformational change and transport a cargo down the gradient (uniporter), or against the gradient driven by the gradient of a co-substrate (indicated in red, symporter and antiporter). (b) Functional background as listed on PlasmoDB for P. falciparum MTP genes [32]. Functional data: MTPs with an annotation supported by functional data, putative function: MTPs with an annotation supported by bioinformatical predictions, unknown function: no functional annotation.
the Plasmodium species, ~140 genes are identified that encode MTPs, with the vast majority to be assessed as carriers [31].
Remarkably, there seems to be only very little redundancy among the transporters. In a systematic study by Kenthirapalan et al., it was shown for around three-quarters of the investigated orphan MTPs that genetic deletion of the gene leads to growth defects at some point during life cycle progression in P. berghei [33]. In accordance with this trend, a recent review pools the phenotypical descriptions after MTP deletions in the current literature and confirms a growth-related role of approx. 78% of Plasmodium’s MTPs during the parasite’s development [30]. Accordingly, a selection of essential MTPs is suggested as potential drug targets due to their exclusive conservation among Plasmodium parasites [31]. Therefore, MTPs are increasingly discussed as rewarding drug-targets, as some candidates show high potency in their safety and efficacy profiles [34]. One example is the P-type ATPase ATP4, which shows susceptibility to a broad range of drugs with a fast killing of blood- stage parasites and with promising candidates in clinicals trials [35, 36]. In contrast, multiple transporters have been identified to induce resistances against antimalarial compounds through mutations in the MTP gene, which confer the ability of drug export [37, 38]. Together, this describes the increasing focus on Plasmodium’s MTPs and their physiological function, which still remains enigmatic for a majority of the proteins.
5 1ST CHAPTER - GENERAL INTRODUCTION
Only about one third of the genes carry functional data to support a characterization. And still, the vast majority of Plasmodium MTP annotations derive mainly from bioinformatical predictions (Fig. 2 b) [39, 40]. Additionally, a categorisation based on structural features of already characterised MTPs allows a classification of even poorly understood proteins into transporter families [41]. However, such annotations should be interpreted with caution, since substrate specificity may be transmitted only by a small region of the protein. Therefore, two structurally very similar MTPs could have entirely different functional features [29].
Despite the description of MTPs as increasingly relevant for the understanding of the parasite’s biology and antimalarial drug therapies, the majority of MTPs still lack robust functional data.
1.4 Towards a functional classification of MTPs
A significant key to understanding the biological role of a transporter is the identification of its cargo. Functional expression in heterologous systems allows for the analysis of trans-membranous transport of isolated transporters. However, such assays have a low throughput which makes previous cargo predictions essential. Therefore, substrate transport via heterologous expression has only been described for very few Plasmodium transporters [30]. Alternatively, in vitro transport assays, for instance on isolated parasites, allow for tracking the transport of substrate-sensing probes. For this purpose, a specific link to the respective protein is needed. This could be either the application of MTP specific drugs or the introduction of mutations in the respective gene, for which an interference with the MTPs transport dynamics can be measured (for instance [42]). Both approaches require an advanced background knowledge on transport features of the MTP and can serve to confirm predictions. However, precise functional hints are not present in many MTPs.
Experimental genetics became a powerful tool to address Plasmodium proteins and to understand the functional role of MTPs. While the first transfections were still relatively challenging compared to other organisms [43], the techniques significantly evolved during the last decades [44, 45], up to the first successful P. falciparum CRISPR/Cas9-mediated genome edits in 2014 [46, 47]. In P. berghei, recombinations showed to have higher efficiencies and to be less time consuming. Additionally, the murine malaria system allows for access to all parasite stages by transmission to the
6 1ST CHAPTER - GENERAL INTRODUCTION
Anopheles vector, while P. falciparum culture is often limited to blood stage cultures. Therefore, the murine malaria system became an advanced and accessible approach to profile Plasmodium’s gene repertoire [48]. Experimental genetics in P. berghei and P. falciparum represent two broad systems which complement each other, and together form a potent tool to characterise Plasmodium’s genome. In the context of MTPs, it facilitates filling the gaps in literature and unravelling potential functions, for instance through the identification of the localisation or phenotyping of MTP-deficient mutants. Still, cross species comparisons might be viewed with caution. Even though major differences between P. berghei and P. falciparum are unlikely for MTPs, some examples have been described [30].
Nevertheless, experimental genetics embody limitations about functional interpretations. But they allow to address aspects which, together with the literature and other techniques can lead towards a better understanding of MTPs.
7 1.5 Aims of the study
Most MTPs have only minor annotations and their functional role remains enigmatic. Therefore, this study addresses a broad selection of MTPs and covers characterisation approaches on a diverse repertoire of proteins that differ in their depth of description in literature, their predicted functions, and determined substrates:
(1) For a small group of transporters, the substrate was confirmed, like for instance for the folate transporter FT2. In the case of folate, an internal de novo synthesis is present as well. Therefore, a functional importance of the transporter might be only of facultative character. In order to unravel a functional role in the parasites cell biology, we study localisation and effects after genomic manipulation of the folate Transporter FT2 .
(2) Depending on the uptake of substrates, MTPs are likely to show host dependent differences, which might correlate with the virulence of the respective Plasmodium sp.. Presence or absence of a particular transporter translates to meaningful differences between the species in the respective biological pathway. To this end, we analyse a putative P. falciparum aminophospholipid transporter, which is absent in rodent malaria parasites.
(3) Research on drug associated MTPs, like the Chloroquine Resistance Transporter (CRT) and ATP4, focuses mainly on the asexual blood stages of P. falciparum. However, a physiological role of the two MTPs in all other stages still remains enigmatic. Here, we identify potential functions beyond blood stages of the well-studied proteins CRT and ATP4 in P. berghei, by analysing the spatio-temporal expression pattern of fluorescently tagged MTPs.
In this study we want to contribute findings towards a functional assignment of MTPs in P. berghei and P. falciparum by the application of experimental genetics in both organisms. As a complement to understanding this increasingly relevant, yet enigmatic protein group, we address the advantages of each of the two Plasmodium systems. All approaches, aim to deliver additional aspects about the MTPs relevance, which could lead to the suggestion of potential new antimalarial drug candidates.
8 2ND CHAPTER - MATERIALS AND METHODS
2nd Chapter: Materials and Methods
2.1 Materials The laboratory work was performed at Humboldt University, Berlin and The Australian National University, Canberra. The Materials listed are a combined collection of both laboratories. Equivalent instruments and material of different producers were only listed once.
2.1.1 Experimental organisms Mouse strain Reference Mus musculus Naval Medical Research Charles River Laboratories (Germany) Institute (NMRI) mice Mus musculus Swiss Webster Charles River Laboratories (Germany) Mus musculus C57BL/6NCrl Charles River Laboratories (Germany)
Mosquito vector Reference Anopheles stephensi Nijmegen (Netherlands) [49]
Microorganisms Reference Plasmodium berghei ANKA Nijmegen (Netherlands) [50] Huh7 human hepatoma cells human hepatoma cells Escherichia coli XL-1 Blue Stratagene (Germany) Escherichia coli PMC103 [51]
This study was carried out in strict accordance with the German ‘Tierschutzgesetz in der Fassung vom 22. Juli 2009’ and the Directive 2010/63/EU of the European Parliament and Council ‘On the protection of animals used for scientific purposes’. The protocol was approved by the ethics committee of the Berlin state authority (‘Landesamt für Gesundheit und Soziales Berlin’, permit number G0294/15). C57BL/6NCrl mice were used for sporozoite infections. All other parasite infections were conducted with SWISS mice.
2.1.2 Plasmodium targeting vectors Vectors Reference pBAT-SIL6 [52] pBAT [52] pGlux6 [53] pCC1 [54] pDC2 [55]
9 2ND CHAPTER - MATERIALS AND METHODS
2.1.3 Enzymes Enzymes Manufacturer Phosphatases and ligases (Kit) Roche (Switzerland) T4 DNA ligase Promega (US) T4 DNA ligase New England Bio Labs (US) Antarctic phosphatase New England Bio Labs (US) S7 Fusion high-fidelity polymerase Biozym (Germany) Taq-DNA polymerase Fermentas (Germany) Q5 high-fidelity polymerase New England Bio Labs (US) Restriction endonucleases New England Bio Labs (US) SYBR Green I Master Mix Thermo Fisher Scientific (US)
2.1.4 Antibodies Anti-P28 antibody (mouse), 1:500 [56] Anti-mouse Alexa Fluor 568-conjugated secondary antibody, 1:1,000 (Thermo Fisher Scientific)
2.1.5 Media, buffers, stock and working solutions Tissue culture water Milli-Q water, autoclaved
Cytomix 120 mM KCl, 0.15 mM CaCl2, 10 mM K2HPO4 pH 7.6, 25 mM HEPES pH 7.6, 2 mM EGTA pH 7.6, 5 mM MgCl2 in tissue culture water adjusted to 7.6 pH, stored at 4°C
Freezing solution Alsever’s Solution 90 ml, Glycerol 10 ml, stored at 4°C
GYT medium 10% (v/v) glycerol, 0.125% (w/v) yeast extract, 0.25% (w/v) tryptone
Heparin/PBS Heparin 40 μl (200 units/ml), PBS 960 μl, stored at 4°C
LB-Agar LB-Medium 1,000 ml, Bacto-Agar 15 g, Solution stored at 4°C
Lysogeny Broth (LB)
Bacto-tryptone 10 g, Bacto-yeast extract 5 g, NaCl 5 g, ddH2O to 1,000 ml, adjust pH 7.5, stored at RT
Lysis buffer 20 mM Tris, 5 mM EDTA, 0.008% saponin (w/v), 0.08% TritonX100 (w/v), adjusted pH 7.5 and autoclaved, stored at RT
10 2ND CHAPTER - MATERIALS AND METHODS
Nycodenz stock KCl 3 mM, EDTA 0.3 mM, Tris pH 7.5, 5 μM, Nycodenz 110.4 g, ddH2O to 400 ml, Solution autoclaved and stored in the dark at 4°C
RPMI complete RMPI with 10 mM glucose, 480 µM hypoxanthine, 20 µg/ml Gentamicin, 0.25% (w/v) ALBUMAX II, 5% (w/v) heat inactivated human serum.
Percoll solution RPMI complete (37°C) 27.1 ml, Percoll (37°C) 14.2 ml, 10x PBS (autoclaved, 37°C) 1.6 ml
Pyrimethamine H2O
Pyrimethamine 0.35 g, DMSO 50 ml, H2O add 5 L, adjust pH 3.5-4.5 and stored in the dark at RT
TAE-buffer (50x) Tris 2 M, Sodium acetate 250 mM, EDTA 0.5 mM, adjust to pH 7.8, stored at RT
Transfection Medium RPMI 1640 160 ml, FCS (origin US) 40 ml, Gentamycin 50-60 μl (40 mg/ml), Solution stored at 4°C
2.1.6 Laboratory equipment Instruments Manufacturer Amaxa Electroporator Amaxa (Germany) Agarose Gel Casting System Easy Cast Thermo Fisher Scientific (US) Acanti J-E Centrifuge Beckman Coulter (US) AxioImager Z2 epifluorescence microscope Zeiss (Germany) AxioCam MR3 camera Zeiss (Germany) Balances Sartorius (Germany) Bench top centrifuge 5424 Eppendorf (Germany) Bench top refrigerated centrifuge 5417 R Eppendorf (Germany) BD FACS – CantoII Flowcytometer BD Bioscience (Germany) BD FACS – Aria cell sorter BD Bioscience (Germany) Binocular MS 5 Leica (Germany) DM 500 microscope Leica (Germany) Electrophoresis power supply Amersham Pharmacia Biotech (UK) Electroporator Genepulser II BioRad (US) Electroporator Genepulser Xcell BioRad (US) Fridge 4°C Liebherr (Germany) Freezer -20°C Liebherr (Germany) Freezer -80°C Heraeus (Germany) Genepulser 2 Bio-Rad (Germany) Gel documentation system Gel Doc XR+ Bio-Rad (Germany) Gilson micropipettes Abimed (Germany) Heating block Eppendorf (Germany) Hemocytometer Marienfeld (Germany) HERAsafe KS12 KS sterile hood Thermo Fisher Scientific (US) Ice machine Ziegra (Germany) Incubator P. falciparum culture Heraeus (Germany) Incubator P. berghei cultures Mytrom (Germany)
11 2ND CHAPTER - MATERIALS AND METHODS
Incubator for mosquitoes Mytrom (Germany) Incubator E. coli plates Memmert (Germany) Incubator and shaker for liquid E. coli culture New Brunswick Scientific (US) Inverted Microscope primo vert Thermo Fisher Scientific (US) Liquid nitrogen container, Arpege 170 Air Liquide (Germany) MACSQuant® benchtop Flowcytometer Miltenyi Biotec (Germany) Magnetic stirrer IKA Labortechnik (Germany) Micro centrifuge Novis (China) Mice facility incubator Ehret (Germany) Mice cages Ehret (Germany) Mosquito cages BioQuip Products Inc (US) Nanodrop ND 1000 peQLab (Germany) PCR Thermocycler, lab cycler Sensequest (Germany) plate reader, synergy HTX Biotek (US) Pipetus Hirschmann Laborgeräte (Germany) pH-meter Mettler Toledo (Germany) Shaker DRS-12 Heraeus (Germany) Sterile bench, Herasafe KS12 Heraeus (Germany) Thermal cycler, MJ Mini, gradient Bio-Rad (Germany) Vortex-Genie2 Scientific Industries (US) Water bath VWR (Germany)
2.1.7 Consumables Consumables Manufacturer 6-well plate Sarstedt (Germany) 8-well glass bottom chamber slide, LabTek Nunc (Germany) 8-well plastic bottom chamber slide, LabTek Nunc (Germany) 24-well plate Sarstedt (Germany) 48-well plate Sarstedt (Germany) Cell culture flask (T-25 and T-75) Sarstedt (Germany) Cell strainer BD Bioscience (US) Cover slips Roth (Germany) Cryo-freezing tubes Greiner (Germany) Electroporation cuvettes (P. berghei) Amaxa (Germany) Electroporation cuvettes (0.2 cm gap) BTX Harvard Aparaturs (US) Eppendorf tubes, 1.5 ml Eppendorf (Germany) Eppendorf tubes, 2.0 ml Eppendorf (Germany) FACS tubes Sarstedt (Germany) Falcon tubes 15 ml and 50 ml Greiner (Germany) Forceps Neolab (Germany) Glass slides, clear Roth (Germany) Glass slides, frosted Roth (Germany) Microplate 96-well, μ-clear (U-bottom) TPP (Switzerland) Microplate 96-well, μ-clear (V-bottom) TPP (Switzerland) Microscope oil Zeiss (Germany) Nature cotton Hartmann (Germany) Needles, sterile Braun (Germany) Pasteur pipettes Kimble (US) PCR tubes Greiner (Germany) Petri dishes Greiner (Germany) Pipette tips Sarstedt (Germany) Round glass cover slips Roth (Germany) Serological pipettes 5 ml, 10 ml and 25 ml Sarstedt (Germany) Sterile filter 0.2 mm and 0.4 mm Sarstedt (Germany)
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Sterile filtration units (500 mL) Sarstedt (Germany) Syringes 1ml and 5 ml Braun (Germany) Tissue culture gas (1% O2, 5% CO2, 94% N2) BOC Gasses and Coregas Transfer pipette 3.5 ml Sarstedt (Germany)
2.1.8 Commercial kits Commercial kits Manufacturer HiFi DNA Assembly Reaction Kit (Gibson) New England Biolabs (US) Pure Link Hi pure plasmid DNA Kit Invitrogen (US) QIAprep Spin Miniprep Kit QIAGEN (Germany) QIAprep Spin Midiprep Kit Q QIAGEN (Germany) QIAamp DNA-Blood Mini Kit QIAGEN (Germany) QIAquick PCR Purification Kit QIAGEN (Germany) QIAquick Gel Extraction Kit QIAGEN (Germany) Rapid DNA Dephos & Ligation Kit Roche (Switzerland) Rapid DNA Ligation Kit Roche (Switzerland) Retroscript Ambion (Germany) RNase-free DNAse set Qiagen (Germany) RNeasy Mini Qiagen (Germany)
2.1.9 Chemicals Reagents Company ß-Mercaptoethanol Sigma (US) 1 kb DNA-Ladder MBI Fermentas, Thermo Fischer (US) 6 x DNA loading dye MBI Fermentas, Thermo Fischer (US) Acetic acid (Glacial) Ajax Finichem (Australia) Agarose Invitrogen (US) Alsever’s solution Sigma (US) AMAXA T cell nucleofector Lonza (Canada) Ammonium acetate Sigma (US) Ammonium persulfate (APS) Roth (Germany) Albumin fraction V Roth (Germany) ALBUMAX II Thermo Fisher Scientific (US) Ampicillin Roth (Germany) Bacto-yeast Roth (Germany) Bacto-tryptone BD Biosciences (US) Cellulose (Carboxymethyl) Sigma (US) Cutsmart Restriction Enzyme Buffer New England BioLabs (US) Chloroform Roth (Germany) Dulbecco's modified Eagle Medium (DMEM) GibcoBRL (Germany) DMSO (Dimethyl sulphoxide) Merck (Germany) dNTP-mix MBI Fermentas (Germany) Ethanol, absolute AppliChem (Germany) Ethanol, 96% Merck (Germany) Ethylene glycol tetra acetic acid (EGTA) Merck (Germany) Ethylenediamine tetra acetic acid (EDTA) Merck (Germany) Ganciclovir Roche (Switzerland) FCS (Fetal Calf Serum), certified (US) Gibco Invitrogen (Germany) FCS Gibco Invitrogen (US) Fibrous Cellulose Powder Sigma (US) Formaldehyde Merck, Darmstadt (Germany) GeneRuler 1kb Plus DNA Ladder MBI Fermentas, Thermo Fischer (US)
13 2ND CHAPTER - MATERIALS AND METHODS
Gentamycin Gibco, Life Technologies (Germany) Giemsa stain VWR (Germany) Glass beads, unwashed Sigma (US) Glucose Merck (Germany) Glutamine Sigma (US) Glycerol Roth (Germany) Glycine Roth (Germany) HEPES (N-2-hydroxyethylpiperazine-N'-2- Sigma (US) ethane sulfonic acid) Heparin Ratiopharm (Germany) Hoechst 33342 Invitrogen (US) HEPES Merck/Calbiochem (Germany) HCl Roth (Germany) Hypoxanthine Sigma (US) Immersion oil Roth (Germany) Isoflurane Baxter (Germany) Isopropanol Roth (Germany) Ketamine/Xylazine hydrochloride Ceva (Germany)/medistar (Germany) L-Glutamine Gibco, Life technologies (Germany) LB-Agar powder (Lennox L Agar) Invitrogen (US) Methanol Thermo Fisher Scientific (US) NaOH Merck (Germany) NAHCO3 Roth (Germany) Nycodenz density gradient medium Axis Shield (Norway) pABA Sigma (US) PBS, tablets Gibco Invitrogen (Germany) Percoll GE Healthcare (US) Penicillin-Streptomycin, liquid Invitrogen (US) Perm-Wash Buffer BD Bioscience (Germany) Potassium chloride Univar (US) Potassium phosphate dibasic trihydrate Merck (Germany) (K2HPO4·3H2O) Potassium phosphate monobasic (KH2PO4) Sigma (US) Pyrimethamine Sigma (US) RPMI 1640-medium GibcoBRL (Germany) Saponin Sigma (US) Sodium chloride Roth (Germany) Sodium acetate Roth (Germany) Sodium bicarbonate Roth (Germany) Sodium hydrogen carbonate Roth (Germany) Sodium hydroxide Merck (Germany) Sorbitol Sigma (US) SOC Outgrowth Medium New England Biolabs (US) SYBR Safe DNA Gel stain Invitrogen (US) Tris (hydoxymethyl)aminomethane (TRIS) Roth (Germany) Trypsin-EDTA Invitrogen (US) Triton X-100 Roth (Germany) Trizol Invitrogen (US) Tween 20 Roth (Germany) WR99210 Jacobus Pharmaceutical (US) Xanthurenic acid Sigma (US)
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2.1.10 Software and databases Adobe Illustrator (Adobe Inc.) Serial Cloner 2.6.1 (SerialBasics) EndNote X9 (Microsoft) SnapGene (GSL Biotech LLC) FlowJo 8.7.4. (Becton, Dickinson & Comp.) GraphPad Prism 7 (GraphPad Software Inc.) ImageJ/Fiji (open source) Microsoft Office 2016 (Microsoft) RStudio (RStudio, PBC)
2.2 Methods 2.2.1 Molecular- and microbiological methods for P. berghei and P. falciparum 2.2.1.1 Polymerase Chain Reaction (PCR)
PCR is applied in various different contexts like molecular cloning or diagnostic PCR. For molecular cloning high fidelity polymerases were used together with specific primers and the reagents listed in Tab. S 1, Tab. S 2 and Tab. 1. For diagnostic PCR used for parasite genotyping or colony PCR, a conventional Taq-polymerase was used. Plasmodium parasites have a (A + T)-rich genome sequence which does often not allow to apply conventional polymerase chain reaction protocols [57]. Therefore, annealing and elongation temperatures were adjusted as listed in Tab. 2 . Additionally, the elongation time was adapted to the according size of the amplified fragment (~1 minute for 1.0 kb). For molecular cloning, the PCR products were purified with a PCR purification kit following the manufacturer’s protocol. Products were stored at -20°C
Tab. 1 PCR component-mix for high fidelity (HF) and Taq-polymerase in P. berghei (P.b.) and P. falciparum (P.f.) P. berghei P. falciparum P. b. & P. f. HF polymerase HF polymerase Taq polymerase component volume (µl) volume (µl) volume (µl) polymerase buffer 5 10 5 dNTPs (10mM) 5 1 1 primers (10 µM) 2 x 1.5 2 x 2.5 1 template (<1000ng) 1 1 1 polymerase 0.2 (0.008 U/µl) 0.5 (0.02 U/µl) 0.25 (0.025 U/µl) water to 50 to 50 to 50
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Tab. 2 Time and temperature settings for P. berghei (P.b.) and P. falciparum (P.f.) polymerase chain reaction for high fidelity and Taq-polymerase. *=optimum temperature can vary and might need adjustment. P. berghei P. falciparum step process temperature time temperature time 1 initial denaturation 94°C 1.5 min 98°C 30 s 2 denaturation 94°C 30 s 98°C 10 s 3 annealing of primers 55°C * 30 s 60°C * 30 s 4 elongation 60°C 1 min 60°C 1 min 5 final elongation 60°C 1 min 72°C 7 min 6 storage 4°C ¥ 4°C ¥
2.2.1.2 Agarose gel electrophoresis
Agarose gel electrophoresis is an electrophoretic separation of mixed DNA fragments based on the size and charge over an agarose gel on which an electric field is applied. The gel was prepared by adding agarose to 1x TAE buffer in a concentration between 0.8% and 1.0% (v/w). The solution was heated in the microwave for 1 minute until agarose was completely dissolved. The solution was allowed to cool down to approx. 50°C, mixed with 10 µg/100 ml SYBR safe and poured in a prepared agarose gel frame with comb to solidify. After 20-30 minutes, the gel was placed in the electrophoresis chamber, covered by TAE buffer, samples mixed with 1 x loading dye and samples and ladders loaded in the wells. Dependent on size of gel and products, the voltage of the electric field was set to 90-110 V and let run for 30-60 minutes. Under UV-light, the DNA bands were imaged for analysis. Depending on the context, bands were cut out and gel purified with the QIAquick Gel Extraction Kit.
2.2.1.3 Digest of DNA with restriction enzymes
Restriction digest was used to cut plasmids and DNA fragments on restriction sites with the respective restriction enzyme, for instance, prior to the ligation of fragments in a plasmid or test digests. The digest was prepared as described in Tab. 3. and dependent on necessity digested at 37°C 15 minutes – 16 hours. For test digests, the digest can directly be applied to a gel. For molecular cloning, purification with the QIAquick PCR purification Kit (Qiagen) is necessary. Digests can be stored on -20°C.
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Tab. 3 Set up of restriction digest for 50 µl.
reagents volume (µl) 10x reaction buffer 5 DNA ~1 µg H2O to 50 restriction enzyme 0.5-1 (5-20 units)
2.2.1.4 Ligation of plasmids and DNA fragments
Ligation describes the fusion of two DNA fragments by a ligase. Dependent on the context, DNA fragments have to be dephosphorylated. Both steps were performed with the Dephos & Ligation Kit (Roche) following the manufacturer’s protocols. The ratio of plasmid to fragment was adjusted 3:1. To increase ligation efficiency, the reaction was incubated overnight at 4°C. Ligation products can be stored at -20°C or directly transformed into competent Escherichia coli.
2.2.1.5 Preparation of electrocompetent PMC103 E. coli
To prepare new stocks of electrocompetent E. coli in the mid-log phase, a single colony of electrocompetent E. coli, which was previously plated out on an antibiotic free LB-agar plate, was picked and inoculated in 50 ml of LB medium. The bacterial culture was incubated overnight at 37°C while shaking at 250 rpm. The next day, two septate flasks of 500 ml prewarmed LB medium were inoculated with 25 ml overnight culture each and incubated and shaken at 250 rpm. Every 20 minutes, the OD600 was measured and blanked against LB medium without bacteria. When the OD600 reached 0.4, the cultures were rapidly transferred on ice and cooled down for 15-30 minutes with occasional swirling. In precooled centrifuge bottles the cultures were centrifuged at 2,500 rpm (precooled Sorvall GSA rotor) for 20 minutes at 4°C. The pellet was resuspended thoroughly in 500 ml ice-cold DI-water. The cells were centrifuged again at 2,500 rpm for 30 minutes at 4°C and resuspended in 250 ml of ice cold 10% glycerol. After centrifugation at 2,500 rpm for 30 minutes at 4°C, the cells were resuspended in 10 ml of ice cold 10% glycerol. After another centrifugation, the cells were resuspended in 1 ml GYT medium. To assess the density of cells in the medium, the OD600 of a 1:100 dilution of the cell suspension was measured and the concentration determined 8 based on the reference, that 2.5 x 10 cells/ml measure 1.0 OD600. The cells were
17 2ND CHAPTER - MATERIALS AND METHODS diluted with GYT to a concentration of 2 x 1010 to 3 x 1010 cells/ml. The diluted cell suspension was shock frozen in 40 µl aliquots and stored at -80°C.
2.2.1.6 Transformation in competent E. coli
For molecular cloning in P. berghei, plasmids where transformed in XL1-Blue competent cells. 35 µl of cells thawed on ice were mixed with 5 µl of the ligation mix and incubated for 10 minutes on ice. Afterwards a 45 second heat shock was performed at 42°C. The cells were cooled another 2 minutes on ice and plated on LB- agar plates with the respective antibiotic. Overnight incubation at 37°C allowed to pick colonies with the transformed plasmid on the next day.
For P. falciparum transformations, electroporation was performed on PMC103 cells [51]. The cells were thawed on ice and 40 µl of cell suspension was gently mixed with 1 – 5 µl of ligation product. The suspension was transferred to a precooled 0.2 cm gap electroporation cuvette without creating bubbles. For the electroporation, the electroporator was set at 1.8 kV, 25 µF and 200 W. The electroporation gives a time constant as a feedback which should be approx. between 4.6 and 4.9 msecs. The suspension was immediately removed from the cuvette, and incubated in 900 µl prewarmed SOC medium for 1 hour at 37°C. Afterward, the SOC medium suspension was diluted 1:100 in SOC medium and plated on LB-agar plates with the respective antibiotic. Overnight incubation at 37°C allowed to pick colonies with the transformed plasmid on the next day.
Colonies were either screened by colony PCR as described above or the plasmid was isolated from bacterial overnight cultures as described below.
2.2.1.7 Plasmid DNA isolation
To isolate the plasmid from E. coli cells, the bacterial culture was expanded after inoculating a 5-10 ml LB-medium culture with antibiotics, by picking cells either from bacterial colonies on LB-agar plates or by transferring a few µl from a previous culture. The inoculated media were incubated in a shaker at 200 rpm overnight. The next day, the cells were harvested by centrifugation and the plasmid was isolated using the QIAprep Spin Miniprep Kit (Qiagen) as described in the manufacturer’s protocol.
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2.2.1.8 Molecular cloning with Gibson assembly
A Gibson assembly is a molecular cloning method, which allows to assemble up to 5 fragments in a plasmid in one single step. Fundamental for this reaction are the homologous overlaps franking the fragments which allow a reassembly based on exonucleases, polymerases and ligases in the master mix. The fragments with the homologous overlaps are designed following the NEBuilder HiFi DNA Assembly’s (NEB) descriptions with primers carrying the overlapping sequences on the 5’ end of the oligomer. The products were amplified by PCR and purified. The assembly reaction was carried out following the manufacturer’s protocol. It is important for the assembly that there is an adjustment of equimolarity for all the used DNA fragments.
2.2.1.9 Isolation of RNA with Trizol and Glycogen
For the extraction of RNA from infected erythrocytes, 1 ml Trizol per 100 µl blood was added and homogenised by passaging through the cannula of a syringe multiple times. The homogenised sample was frozen overnight at -80°C and defrosted the next day. 500 µl of the solution was transferred in an RNase free tube and 200 µl chloroform was added. The tube was inverted 15 times and allowed to settle for 3 minutes at room temperature. Afterwards, the solution was centrifuged at >13.000 rpm for 15 minutes at 4°C. 100-120 µl of the supernatant’s upper layer was transferred in a new tube. To precipitate the RNA, 500 µl of isopropanol, 120 µl of 5 M ammonium acetate, and 10 µl of ice-cold glycogen were mixed and incubated at -20°C for at least 20 minutes. Afterwards, the solution was again centrifuged at >13.000 rpm for 15 minutes at 4°C and the supernatant was discarded. To wash the RNA, the pellet was resuspended in 1 ml of ice-cold 70% ethanol by vortexing and centrifuged at >13.000 rpm for 10 minutes at 4°C. The supernatant was aspirated and the pellet allowed to dry for 10 minutes without over-drying it. Finally, 50 µl of RNase/DNase-free water was used to re-dissolve the pellet at 65°C for 5 minutes. The RNA concentration was measured with a 1:100 diluted sample at the 260/280 ratio in a NanoDrop. The samples can be stored at -20°C but are the most stable after being converted to cDNA.
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2.2.1.10 Sanger Sequencing
DNA sequencing was performed with 13 µl of DNA (600 - 1500 ng) together with 1 µl of the respective sequencing primer (10 µM) via agrf (Australia) or LPC Genomics (Germany)
2.2.1.11 Isolation of genomic DNA
The genomic DNA isolation from isolated parasites was performed with a QIAamp DNA Blood and tissue kit (Qiagen) according to the manufacturer’s protocol and stored at -20°C.
2.2.1.12 DNA precipitation
The DNA, for instance a linearized plasmid, was mixed with 1 ml ice cold 99% ethanol with 40 µl 3 M Na acetate for 1 hour at -80°C. After centrifugation for 15-60 minutes at 4°C, the supernatant was discarded and the pellet resuspended in 500 µl 70% ethanol. After additional centrifugation for 10 minutes, the supernatant was removed completely and the pellet was allowed to dry for 5-10 minutes (without over- drying). Afterwards, the DNA pellet was resuspended in 20 µl DI-water and dissolved in a shaker for 10-30 minutes at 37°C.
2.2.1.13 Complementary DNA (cDNA) conversion by reverse transcription.
The RNA was converted to cDNA in a reverse transcription reaction PCR by using the Retroscript Kit following the manufacturer’s protocol.
2.2.1.14 Quantitative real-time PCR (qPCR)
qPCR is a quantitative PCR reaction based on cDNA converted from the transcript, which allows to quantify the transcription of a gene of interest compared to a house keeping gene with stable expression. Specific primers with an exon-overlap amplify a product from a cDNA template. The product intercalates a fluorescent dye. Real-time measured fluorescence signal during the PCR cycles, serves as the feedback for the quantification of the cDNA/transcript. Per reaction well, 5 µl of 1:30
20 2ND CHAPTER - MATERIALS AND METHODS diluted cDNA was applied and 7.5 µl of a master mix was added (Tab. 4). The reaction was performed with the time and temperature program listed in Tab. 5.
The relative expression of the gene of interest in relation to the housekeeping [Ct sample-Ct housekeeping gene] gene was calculated by using the delta Ct-method (R=2 ).
Tab. 4 Composition of qPCR master mix
reagent volume per sample (µl) SYBR Green I Master Mix 6.25 Fwd primer (100 µM) 0.025 Rev primer (100 µM) 0.025 water 1.2 total 7.5
Tab. 5 qPCR protocol with time and temperature steps repeat temperature time 1x 95°C 15 min 40x 95°C 15 sec 55°C 15 sec 60°C 45 sec data collection 1x 95°C 15 sec 60°C 20 min 95°C 15 sec melting curve
2.2.2 Live fluorescence microscopy
All fluorescence imaging was performed on an AxioImager Z2 epifluorescence microscope equipped with an AxioCam MR3 camera. Parasite stages were prepared for live microscopy as described in the life cycle analysis below. To avoid bleaching, previous light exposure was avoided and parasites were imaged only once. Appropriate exposure times were adjusted for each fluorescence channels on a test parasite which was not recorded. To get the parasite in focus, differential interference contrast microscopy (DIC) was used. The fluorescence channel order for imaging was mCherry, GFP, DIC, DAPI. Live imaging was performed rapidly with fresh samples.
In measurements for spatio-temporal MTP::mCherry signal-quantification the exposure-time settings were kept identical measured parasite stages. Identical settings were applied to a wild type (WT) control to quantify autofluorescence. The signal output of the fluorescence channels was analysed from zvi-files using the software imageJ (Fiji). Every pixel of the image is associated with a signal intensity,
21 2ND CHAPTER - MATERIALS AND METHODS which was quantified using Fiji’s measurement function on the area of the whole parasite. Cytoplasmic expressed GFP allows to determine the total area of the respective parasite stages. The used digital output for one measurement was the sum of raw intensity of all pixels integrated to the measured area (integrated densityparasite), as well as area and mean pixel intensity. By measuring three additional fields of the same size outside of the infected erythrocyte, background noise was identified and subtracted from the parasite measurements and transformed into integrated density/µm2 using the following equation:
Integrated density Integrated density − (mean pixel intensity x area ) = µm area
According to this, the integrated density/µm2 reflects the signal from MTP::mCherry only. The consideration of signal per µm2 allows a general analysis of the expression signal independent of a potential spatial overrepresentation by the localisation in a specific cellular compartment. Identical measurements were performed on wild type parasites and mean autofluorescence subtracted from the mean parasite signal.
2.2.3 Transfections of recombinant P. berghei lines and phenotyping 2.2.3.1 Generation of recombinant P. berghei strain ANKA parasites lines
For endogenous tagging of FT2 (PBANKA_0931500) constructs for single homologous recombination were designed (comparable to Fig. S 3 a). With the forward and reverse primers in Tab. S 1 the carboxy-terminal sequence upstream of the stop codon was amplified by PCR from P. berghei strain ANKA genomic DNA and inserted in a pBAT vector [52] after previous restriction enzyme digestion with EcoRI and HpaI of both, PCR product and vector. After ligation, confirmation of integration and preparation for transfection were performed as described below.
For the following introduction of the api-GFP marker, the promotor of the heat shock protein 70 (HSP70, PBANKA_0711900) which drives a high-level expression of cytoplasmic GFP, was cut out of the pBAT plasmid by restriction enzyme digestion using PvuII and PshAI (Fig. S 3 a). Later on, it was replaced by ligation with the PvuII
22 2ND CHAPTER - MATERIALS AND METHODS and PshAI digested promoter and amino-terminal sequence of the 20 kDa chaperonin gene (CPN20, PBANKA_1347800) produced with forward and reverse primers from Tab. S 1 on genomic DNA. Afterwards, confirmation of a correct integration and preparation for transfection were performed as described above.
The P. berghei strain ANKA FT2 deletion was a targeted by a pBAT vector with 5’ and 3’ flanking regions of FT2. The flanking regions were amplified by PCR with the respective Primers in Tab. S 1 from P. berghei strain ANKA genomic DNA and digested with the restriction enzymes SacII and PvuII for the 5’ flanking region and XhoI and KpnI for the 3’ flanking region, respectively. In two steps, the flanking regions were ligated in the SacII and PvuII or XhoI and KpnI digested pBAT vector. After the confirmation of a correct integration, the plasmids were prepared for transfection as described above.
– For the generation of the ft2comp parasite line, pyrimethamine-resistant ft2 parasites were exposed to an in vivo negative selection with 5-fluorocytosine, as previously described [58]. The ft2– parasites that had lost their drug-selectable cassette through homologous recombination were isolated by limiting dilution cloning. For this purpose, mice were infected with a calculated dose of one parasite per injected volume. The gained strain was used as the recipient strain during transfection with the FT2 complementation construct. This construct was generated by insertion of the CPN20 promoter sequence into the pBAT-SIL6 vector using SacII and EcoRI, followed by the FT2 coding sequence which was cloned in frame with mCherry-3xMyc using EcoRI and HpaI. The cytoplasmic GFP expression cassette was removed by restriction digest with PvuII and SfoI and subsequent religation. The plasmid was linearized with AhdI and ApaLI prior to transfection.
The ft2over parasite line was generated under an identical design as the ft2comp parasite line but on P. berghei ANKA wild type parasites instead of ft2– parasites. Together with the expression of FT2 from the unchanged wild type locus, an additional ectopic FT2 expression in SIL6 leads to an overexpression of the protein in the parasite line.
The constructs for endogenous tagging of ATP4 (PBANKA_0610400) and CRT (PBANKA_1219500) were generated in P. berghei strain ANKA parasites through a gene replacement strategy via double crossover homologous recombination (Fig. 17 a and Fig. 21 a, equivalent to [33]). The 3’ and CT flanking regions of the gene were
23 2ND CHAPTER - MATERIALS AND METHODS amplified by PCR from P. berghei strain ANKA genomic DNA using the respective forward and reverse primers from Tab. S 1. The gained products were digested with SacII and EcoRI for the CT flanking region and KpnI and AvrII for the 3’ flanking region. The digested products were ligated in a pBAT-SIL6 vector harbouring a mCherry- 3xMyc-tag [52], which has previously been digested with the respective restrictions site of the PCR product and purified. The two flanking regions were inserted in two ligation steps. Intermediate plasmids were transformed in chemically competent XL1-Blue E. coli in which they were selected against an ampicillin resistance transmitted by the transformed pBAT-SIL6 vector. Ligated plasmids were isolated with plasmid minipreps and the correct integration of the 3’ or CT flanking region was confirmed by sanger sequencing. Additionally, a diagnostic digest was performed with the respective restriction enzymes of the fragment. The final plasmid resulted in an in-frame fusion of the CT coding regions with the mCherry-3xMyc tag. Confirmed plasmids were scaled up in E. coli cultures, isolated in midiprep kits and linearized with SacI and SalI. The linearized plasmid was approved by agarose gel electrophoresis and precipitated. Afterwards the plasmid was prepared for transfection as described below.
2.2.3.2 Transfection of P. berghei schizonts
For P. berghei transfection, the linearized and precipitated vector is transfected in P. berghei schizonts gained from a previous schizont culture as described below. The gained schizonts were pelleted by pulse spinning for 15 seconds. Then 5-10 µg of the prepared plasmid were mixed with 100 µl of human T cell nucleofector AMAXATM. Afterwards, the suspension was added to the pelleted schizonts and carefully transferred to an electroporation cuvette without creating bubbles between the electrodes. The cuvette was electroporated using the U-033 function of an AMAXATM gene pulser. Immediately, the cuvette was taken from the pulser, 50 µl of T-medium (transfection medium) were added and injected intravenously in a naïve mouse. The next day, the mouse was fed with pyrimethamine (70 ng/µl, pH 3.5-3.0) supplemented water. Six days after the parasite infection, the blood from a mouse with a parasitemia >1% was preserved in cryo-stabilates.
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2.2.3.3 P. berghei schizont culture
Blood extracted from an infected mouse contains almost no schizonts. Therefore, an overnight schizont culture allows to enrich this parasite stage from infected mouse blood: a naïve mouse was infected with 107 blood stage parasites to get a parasitemia of 3-5%. The blood extracted from the mouse was added to 10 ml of prewarmed T-medium with 250 µl 1% heparin in PBS and centrifuged for 8 minutes at 1,000 rpm without break. The supernatant was removed and the pellet resuspended in 5 ml of prewarmed T-medium, and pipetted on the bottom of a prewarmed conical flask with 250 ml of prewarmed T-medium underneath the T-medium layer. The opening of the conical flask sealed with cotton wool and incubated at 37°C overnight, aerated with 10% O2, 5% CO2 and 85% N2 for 18-20 hours. On the next day, a Giemsa smear was prepared to assess the developmental stage of the schizonts. When the parasites were ready, 10 ml of 55% Nycodenz in PBS was pipetted under 35 ml of the overnight culture and centrifuged at 1,000 rpm for 25 minutes without break. Schizont were aspirated from a greyish-brown layer above the Nycodenz layer and centrifuged at 1,500 rpm for 10 minutes with break. The supernatant was removed and the pellet resuspended in 1 ml T-medium. The schizont-rich suspension was ready for transfection.
2.2.3.4 Selection of isogenic P. berghei cell line
Depending on the design, the recombination includes the integration of a GFP cassette expressed under the promoter of HSP70, which leads to recombinant parasites expressing a high signal of cytoplasmic GFP. After the confirmation of GFP positive parasites in the parental population by fluorescence microscope, infected blood with a parasitemia between 0.1% and 1% was prepared for the flow cytometry isolation of the recombinant parasites.
One drop of blood from tail puncture was collected and mixed immediately with 1 ml of Alsever’s solution. The solution was applied to a 30 µm Cell Trics filter to separate agglutinations. In the Flow Cytometry Core Facility at the Deutsches Rheuma-Forschungszentrum Berlin the sample was sorted at a FACS Diva as described in [59] by an excitation at 488 nm with a detection band pass filter of 530/30 nm and maximum sensitivity set at the photomultiplier tube voltage. After gating FSC and SSC to single cell erythrocytes, 100 GFP positive cells were sorted into a tube
25 2ND CHAPTER - MATERIALS AND METHODS with 200 µl RPMI supplemented with 20% FCS. This suspension was intravenously injected in a naïve mouse, from which blood was extracted and cryo-preserved with a parasitemia >1%.
2.2.3.5 Cryopreservation of mouse blood infected with P. berghei
To preserve parasite strains as cryo-stabilates, 100 µl of P. berghei infected blood, which was directly extracted from a mouse, was mixed with 200 µl of a freezing solution and frozen immediately after at -80°C.
2.2.3.6 Isolation of P. berghei from infected mouse blood
Blood from a mouse with a parasitemia >1% was extracted and applied to an PBS equilibrated column with a cotton filter, cellulose and glass beads. The blood on the column was topped up with 14 ml PBS and the flow through collected in a 15 ml falcon tube. The suspension was centrifuged at 1,500 rpm for 8 minutes and resuspended in 14 ml 0.2% saponin in PBS. The lysate was centrifuge again at 2,800 rpm for 8 minutes. The pellet was transferred with PBS in a 1.5 ml tube and washed with PBS by centrifugation at 7,000 rpm for 3 minutes. The pellet was then resuspended in 200 µl PBS and stored at -20°C.
2.2.4 P. berghei life cycle analysis 2.2.4.1 Giemsa-stain of thin blood smears
To monitor a mouse infected with P. berghei, a Giemsa stain was performed. By tail puncture a drop of blood was put on a glass slide. With a slow and constant movement, the drop was smeared to a thin blood smear with a second glass slide. After air drying and short fixation in 100% methanol for 10-30 seconds, the slide was stained in a 10% Giemsa solution for 5-10 minutes. After washing the slide under running water, the slide was dried and examined with a bright-field microscope under the 1,000-fold magnification with immersion oil. Intracellular parasites are recognisable by the deep blue Giemsa stain, which dyes the phosphate groups of DNA. To assess the infection, developmental stages of the parasites were observed and the parasitemia was determined (the percentage of P. berghei infected erythrocytes among all erythrocytes).
26 2ND CHAPTER - MATERIALS AND METHODS
2.2.4.2 Asexual blood stage development
For an analysis of the asexual blood stage development of a parasite strain, mixed infections of wild type strain and strain of interest were analysed by an intravital competition assay [60]. The two lines were expressing different cytosolic fluorescents (for instance mCherry expressing Berred wild type and GFP expressing knock out line) and 500 parasites of each line co-infected intravenously into a naïve mouse. The parasitemia was analysed daily by flow cytometry, by gating for infecting erythrocytes and a count of the respective fluorescent colour. Seven days after the infection, 1,000 mixed parasites were transferred into a naïve mouse and the parasitemia was determined on the day of blood passage.
For live microscopy of the blood stages, 107 blood stage parasites were intravenously injected in a naïve mouse. Three to five days after infection, a drop of fresh blood from tail puncture was put on a glass slide and gently mixed with 2-5 µl of RMPI with 1:500 Hoechst. Gently, the suspension was covered with a coverslip and allowed to settle in the dark for 10 minutes. Live microscopy was performed to image from ring stages until gametocytes.
For imaging of fixed cells, 5 µl of infected blood from tail puncture was mixed with 255 µl RPMI. 20 µl of the solution were transferred to poly-L-lysin coated cover slips. The cells were allowed to settle for 10 minutes. Afterwards, the coverslips were fixed for 20 minutes in a well with 300 µl of 4% PFA.
2.2.4.3 Sexual blood stage development
To track the development of sexual stages in a parasite strains, aspects like conversion-rate to gametocytes and quantification of exflagellation-efficiency were inspected. To access gametocytes, 107 blood stage parasites were intravenously injected into a naïve mouse. Three to five days after the infection, a drop of fresh blood from tail puncture was taken and put on a glass slide. For a determination of gametocyte conversion rate, the drop was smeared to a thin blood smear and Giemsa stained as described above. To assess the conversion of asexual blood stages to gametocytes, approx. 2,000 infected erythrocytes were counted under a bright field microscope (1,000-fold magnification with immersion oil) and the percentage of gametocytes per infected erythrocyte was calculated.
27 2ND CHAPTER - MATERIALS AND METHODS
Exflagellation-efficiency was addressed by mixing 5 µl of the blood from tail puncture with 125 µl RPMI supplemented with 50 µm XA. The suspension was transferred to a Neubauer chamber and incubated at 20°C. 10-18 minutes after activation the exflagellation-centres were quantified under a bright field microscope at 400-fold magnification.
2.2.4.4 Ookinete formation culture
For in vitro fertilisation and ookinete conversion, 107 blood stage parasites were intravenously injected into a naïve mouse and 10 μl of infected blood from day 3 after infection were cultivated overnight in 90 μl of RPMI 1640 medium supplemented with
10% fetal calf serum, 50 μM xanthurenic acid (XA) and 0.85 g/L NaHCO3 (pH 8.0) at 20°C. Ookinete cultures were stained live with a mouse anti-P28 antibody (1:500) [61] in combination with an Alexa Fluor 568-conugated secondary antibody (1:1,000) and imaged by fluorescence microscopy. Ookinete conversion was expressed as the percentage of fully developed ookinetes among P28-positive parasites.
2.2.4.5 Breeding of Anopheles stephensi mosquitoes
The breeding of Anopheles stephensi mosquitoes was done in an insectarium with 28°C, 75% humidity and dark/light rhythms of 10 hours/14 hours. The adult imagoes were held in cages and fed with 10% sucrose solution complemented with pABA and a breeding solution of 1% sea salt in DI-water. Larvae and eggs were held in trays with breeding solution. Pupae were collected every day and transferred to the mosquito cages for hedging. To induce female imagoes to lay eggs they were allowed to feed on an uninfected and anaesthetised (80-100 µl ketamine/xylazine hydrochloride) mouse for 30-60 minutes.
2.2.4.6 Transmission of blood stage parasites to Anopheles stephensi mosquitoes
107 blood stage parasites were intravenously injected in a naïve mouse. After day three post infection, an exflagellation assay was performed as described above. If >3 exflagellation centres per field were observed, the feeding was performed. In advance, mosquitoes were starved for 6-10 hours. The mouse was then anaesthetised with 80–100 µl ketamine/xylazine hydrochloride and mosquitoes were allowed to feed
28 2ND CHAPTER - MATERIALS AND METHODS on it for 30-60 minutes. Afterwards, the mouse was sacrificed, and the mosquitoes were kept at 20°C with a daily supply of feeding solutions as described above.
2.2.4.7 Analysis of parasite-infected mosquito midguts
The developing oocyst stages can be monitored during two to three weeks after the transmission to the mosquito. Therefore, mosquitoes were immobilised by being cooled on ice for 10 minutes and being briefly sprayed with ethanol and washed in PBS. During dissection, they were kept in chilled RPMI with 3% BSA. For a potential phenotypical analysis, the midgut was dissected out of the mosquito’s abdomen and placed on a glass slide with 1:1,000 Hoechst in PBS. It was allowed to incubate in the dark for 10 minutes. Afterwards fluorescence microscopy was performed as describe below. To image a whole midgut for an oocyst-count, the midguts were imaged under 25-fold magnification usually at day 10 or day 14 after transmission to the mosquito. For the imaging of oocysts, the upper surface of the midgut was focused with a 640- fold magnification and individual oocysts were imaged.
For the oocyst’s morphological analysis, the live imaged oocysts were captured at their maximal expansion, indicated by cytosolic GFP. The transverse area was digitally measured using ImageJ and the volume was derived from the area using the following equation:
4 A . V = ∗ π ∗ A ∗ 3 π Additionally, morphological features of the oocysts, like vesiculation or sporoblast formation were quantified. The interpretation of a sporulating oocysts, was an oocyst showing typical feature of sporozoite formation. For instance, partly or completely formed sporozoites with aligned DNA around a sporoblast. Depending on the focus, sporozoites only and not the sporoblast were visible. Oocyst showing internal vesicles with a rough surface and displaced cytosol were determined as abnormal. Fully vesiculated oocysts were also included. Oocysts with internal vesicles with a smooth surface were not included to this definition.
29 2ND CHAPTER - MATERIALS AND METHODS
2.2.4.8 Sporozoite extraction of mosquitoes’ salivary glands
In order to access and quantify sporozoites in the mosquitoes’ salivary glands, the mosquitoes were prepared for dissection as described above. The glands were isolated by dissection from the mosquito’s thorax and collected in RPMI with 3% BSA on ice. Afterwards, the glands were grinded until complete homogenisation for at least 10 minutes on ice and then centrifuged for 3 minutes at 10,000 rpm at 4°C. The pellet was resuspended in 300 µl RPMI with 3% BSA and permanently kept on ice. In a 1:10 dilution, the sporozoites were quantified in a Neubauer chamber. For microscopy of the isolated sporozoites, a few microliters of the sporozoite suspension were mixed with Hoechst (1:1,000) and put on a glass slide with a cover slip. After letting them settle for 10 minutes in the dark, fluorescence microscopy was performed under 640- fold magnification.
2.2.4.9 Transmission of sporozoites to naïve mice
For natural transmission to naïve mice, mosquitoes were starved for 6-10 hours before the transmission experiment. A naïve C57BL/6NCrl mouse was anaesthetised and the starved mosquitoes were allowed to feed on the mouse for 15-20 minutes. Afterwards the mouse was put back in its cage and observed during recovery from anaesthesia. The next days, blood of the mouse was checked by tail puncture and controlled on a thin blood smear or by flow cytometry.
Alternatively, 10,000 sporozoites can be intravenously injected into naïve mice. Blood stages were expected to appear at day 3 after infection.
2.2.4.10 In vitro infection of hepatoma cells (Huh7) with sporozoites
Hepatoma cells were continuously cultured in DMEM complemented with 10%
FCS and 2% Penicillin-Streptomycin at 37°C in a gas composition with 5% CO2. A day before the infection experiment 30,000 Huh7 cells/well were seeded on a plastic or glass LabTek slide in 300 µl DMEM complemented medium. The day after, the cells were washed once with DMEM complemented medium. 100 µl of DMEM complemented medium with 10,000 sporozoites/well was applied to the wells and incubated one hour at room temperature to allow the sporozoites to settle down. Afterwards, the plates were placed in the incubator. This is the time point where the
30 2ND CHAPTER - MATERIALS AND METHODS infection experiment started. Two hours later, the medium was exchanged. At 24 hours, 48 hours and 69 hours after infection the medium was exchanged. For live imaging, the medium and the frame of the LabTek slides were removed, a drop of DMEM complemented medium with 1:1,000 Hoechst was applied and covered with a cover slip. After 10 minutes of incubation at room temperature, the cells were imaged under the 640-fold magnification of the fluorescence microscope.
Morphological features like size and volume were measured as described above for oocysts.
2.2.5 Transfection of recombinant P. falciparum 3D7 lines 2.2.5.1 Generation of recombinant P. falciparum 3D7 lines
The construct for episomal tagging of FT2 (Pf3D7_1116500) was generated by using a pGLUX-6 vector [53]. The FT2 gene without stop codon was amplified by PCR from Pf3D7 genomic DNA using forward and reverse primers from Tab. S 2. The product was digested by XhoI and KpnI and ligated with the XhoI and KpnI digested pGlux-6 vector as described above. The Ligation was transformed in PMC103 electrocompetent cells and plasmid mini-preparations were isolated. The ligated vector was checked by Sanger sequencing for the correct integration and prepared for transfection.
The Pf3D7 deletion of FT2 was targeted with a pCC1 [54] vector with 5’ and 3’ homologous regions for double homologous recombination. The 5’ and 3’ homologous regions were amplified by PCR with forward and reverse primers designed with 5’ homologous overlaps for the Gibson assembly on Pf3D7 genomic DNA (Tab. S 2). The pCC1 vector was digested with EcoRI, AvrII, SpeI and SacII to cut out the drug cassette and assembled with the 5’ and 3’ homologous regions as described in the manufacturers protocol of the Gibson assembly kit. Equimolarity was adjusted for all the fragments of the assembly. After transformation and plasmid mini preparation, the ligated plasmid was sequenced by Sanger sequencing.
For a CRISPR/Cas9-mediated disruption of FT2 and Pf3D7_1468600 a pDC2 vector [55] was used to induce a double strand break and allow double homologous recombination. For the amplifications of the 5’ and 3’ homologous regions by PCR, forward and reverse primers from Tab. S 2 were designed with an overhang of homologous overlaps for a reassembly during the Gibson assembly. The pDC2 vector 31 2ND CHAPTER - MATERIALS AND METHODS was digested with ApaI and EcoRI and equimolarity was adjusted with the amplified 5’ and 3’ homologous regions. The assembly was performed as described in the manufacturers protocol of the Gibson assembly kit. The ligation product was transformed in PMC 103 cells and a plasmid was isolated by a mini preparation. The ligated vector was checked by Sanger sequencing for correct integration.
2.2.5.2 Maxi preparation of DNA
P. falciparum transfections require approx. 100 µg of DNA. To scale up the concentrations and quantities of a plasmid, an adjusted maxi preparation (PureLinkTM HiPure plasmid DNA purification kit) of the required plasmid needs to be performed in advance. A 500 ml bacterial overnight culture was incubated in LB medium with antibiotics for 10-16 hours. The plasmid carrying bacteria were either picked from viable cultures or bacterial cryo-stabilates. For harvesting the cells from the culture, the medium was split into two appropriate centrifugation containers and centrifuged for 10 minutes at 4,300*g with a Sorvall SS-34 rotor. The clear supernatant was removed and 30 ml of the purification kits resuspension buffer (R3) containing RNAse A was added and mixed until the complete resuspension of the pellet. Afterwards, 30 ml of lysis buffer (L7) was added, mixed gently by multiple inversions and incubated for 5 minutes. 30 ml of precipitation buffer (N3) was added to the lysate, immediately mixed by multiple inversions and centrifuged for 10 minutes at 12,000 rpm. Previous to centrifugation, the columns of the purification kit were equilibrated with equilibration buffer (EQ). The supernatant of the centrifuged lysate was applied to the column and allowed to drain by gravity flow. The DNA binds the membranes of the column, which was afterwards washed with 60 ml washing buffer (W8, drains by gravity flow). After the completed wash, 15 ml elution buffer (E4) was added to the column to elute the DNA from the membrane. The flow through was caught up in an autoclaved centrifugation tube, 10.5 ml of isopropanol added, mixed and stored at -20°C for 30 minutes. In a precooled centrifuge, the elution was pelleted for 30 minutes at 11,500 rpm. The very small, white to slightly transparent pellet is the purified DNA. Carefully, the supernatant needed to be removed without resuspending the pellet. The pellet was additionally washed with ice cold 99% ethanol and spun at 4°C for 5 minutes at 15,000 rpm. To remove the ethanol completely, the supernatant was discarded and the last millilitres removed with a fine pipette without toughing the pellet. Afterwards, the pellet
32 2ND CHAPTER - MATERIALS AND METHODS was air dried in the biosafety cabinet for 10-15 minutes, without over-drying the pellet. TE elution buffer was prewarmed at 37°C and the pellet was resuspended. Up- and down pipetting for at least 30 seconds increases the elution efficiency. After determining the concentration, the plasmid was stored at -20°C.
2.2.5.3 Preparation of plasmid for transfection
Two days before the transfection, the plasmid was precipitated. Therefore, 100 µg of DNA was adjusted to 100 µl with DI-water and 10 µl of 3 M NaOAc (pH 5.2) was added without touching the DNA solution with the pipet tip. Additionally, 250 µl of 99% ethanol was added and the precipitation was mixed and stored overnight at -20°C. The next day, precipitation was centrifuged at 13,000 rpm for 20 minutes at 4°C. The supernatant was discarded and the pellet was washed by adding 1 ml of 70% ethanol (v/v) followed by centrifugation at 13,000 rpm for 5 minutes at 4°C. The supernatant was removed, and the opened tube was dried in the biosafety cabinet for 5-10 minutes without over-drying the pellet. 15 µl of prewarmed TE buffer was added and mixed. The resuspension was incubated for further 15 minutes at 37°C. Afterwards, the DNA was stored at 4°C overnight to make sure that the pellet completely dissolves.
2.2.5.4 Transfection of P. falciparum rings stages
Parasites were prepared for transfection by keeping a stable culture on fresh blood at least a few days previous to transfection. At least 5 ml of culture needs to be prepared for each transfection. Since ring stages are needed to ensure an uptake of DNA, the culture was synchronised by sorbitol synchronisation (as described below) two days in advance. The day before the transfection, the synchronised parasite culture was assessed by a Giemsa stain. To maintain a culture with high ring stage parasitemia on the day of transfection, the culture should harbour ~2% mature trophozoites on the previous day.
On the day of transfection, the cultured parasitemia was determined and should have >5% ring stages. The DNA precipitation from the previous day was resuspended at 37°C for at least 5 minutes and 15 µl of DNA mixed with 385 µl prewarmed, filtered and sterilised cytomix and kept at 37°C until transfection. More than 5 ml of the culture per planned transfection was centrifuged in a 10 ml tube at 1,500 rpm for 4 minutes.
33 2ND CHAPTER - MATERIALS AND METHODS
The supernatant was discarded and 200 µl of the cultured parasites pellet was gently mixed with 400 µl of DNA in cytomix. This mix was pipetted without bubbles to an electroporation cuvette with a 0.2 cm electrode gap and the pulse electroporated by a BioRad Genepulser Xcell under the parameters in Tab. 6. The electroporator gives as a feedback the respective time constant which has an optimum between 7.5-12 msec. Transfection within this optimum were immediately and gently resuspended in a prewarmed dish with RMPI 1640 with supplements and 3% haematocrit and the cuvette was gently washed with culture medium. Afterwards, the culture was swirled and cultured as mentioned above. 4-6 hours after transfection, the supernatant of the culture was replaced by fresh medium, and the respective antibiotic for selection was added in the required concentration (for instance WR99210 2-10 nM).
Tab. 6 Parameter settings for P. falciparum electroporation in BioRad Genepulser Xcell
parameter setting voltage 0.31 kV capacity 950 µF resistance none
The following 5 days, the medium was replaced with prewarmed RMPI 1640 medium with supplements and the respective antibiotic without taking up RBCs. On day two after transfection, the culture was assessed and the parasitemia was determined. No rings but extracellular trophozoites are expected. Day three after transfection, the culture is again assessed to avoid overgrowth. In case of overgrowth, the drug selection might not have worked. Day 5-12 after transfection, the RPMI 1649 medium with complements and antibiotic was replaced without taking up erythrocytes at least three times a week. Once a week 100 µl of fresh uninfected erythrocytes was added. Additionally to the media changes, 14 days after transfection parasitemia was monitored by Giemsa stains and checked for intracellular parasites. Once parasites were visible, the culture was split and expanded as soon as the parasitemia reached 1%. As soon as possible cryo-stabilates with >5% ring stages were cryopreserved as described above. Dependent on the recombination approach, parasites can be tested immediately by diagnostic PCR for integration of the aimed modification. For conventional recombination by double homologous recombination by pCC1 plasmids, the parasites have to undergo drug cycling.
34 2ND CHAPTER - MATERIALS AND METHODS
Drug cycling describes swapping between on- and off-drug-pressure in multiple cycles. Through that, parasites with an episomal expression of the respective drug resistance will lose their plasmid during the absence of the drug pressure and get eliminated in the following on-drug cycle. Only recombined parasites harbouring the integrated resistance cassette together with the aimed endogenous recombination will remain in the culture. Therefore, cultured populations from new transfections were cultured for three weeks without the presence of the antibiotic. After three weeks, the antibiotic was added in the previous concentrations on a culture with at least 2-3% parasitemia. The next days, the death of the parasites was monitored daily and split when needed. As soon as ring stages were present in the Giemsa smears, the culture was expanded and cryo-stabilates were frozen with high ring stage parasitemia. For the second cycle of drug cycling, the drug-pressure was again removed from the established culture, and cultured for the following 3 weeks without antibiotics. As for the first drug cycle, the cultures were monitored, expanded and cryo-stabilates were prepared. Positive selection, in which parasites with an endogenous integration of the drug cassette were selected, was succeeded by negative selection.
At this point, the culture still harbours parasites in which the drug-cassette for positive selection was integrated via single crossover homologous recombination in the genome. Therefore, the gene is technically only interrupted and not deleted. To overcome this, negative selection was performed by an E. coli cytosine deaminase (CD) which, in case of single homologous recombination, gets integrated with the backbone of the vector. Expression of CD led to a conversion of the non-toxic 5- fluorocytosine (5-FC) into 5-fluoro uracil (5-FU) which inhibits RNA- and thymidylate synthesis [62]. Negative selection eliminates pCC1 backbone-integrated parasites, and selects the rare double homologous recombination events with the desired deletion of the gene. Initially, the parasite line coming from the second positive selection cycle was sorbitol-synchronised. The following days, the parasitemia was monitored and adjusted to 2-4% ring stages and 231 nM 5-Fluorocytosine was applied to the culture, while maintaining the previous concentration of the drug for positive selection. On the following three days, the culture was monitored and medium with 5- Fluorocytosine and the drug from positive selection was exchanged without taking up erythrocytes. Day four onwards, cultures were monitored as before with both drugs three days a week until parasites appeared. After a stable culture was established,
35 2ND CHAPTER - MATERIALS AND METHODS cryo-stabilates were prepared. The parasites can be tested by diagnostic PCR for integration of the aimed modification.
2.2.5.5 Diagnostic PCR for screening of recombinant P. falciparum
To diagnose the correct genomic recombination, a recombination specific PCR was performed on genomic DNA of the transfection-obtained strain. Parasites were isolated from 2-5 ml of the culture via saponin lysis described below and genomic DNA was extracted using a QIAGEN DNeasyÒ Blood & Tissue Kit according to the manufacturer’s protocol and stored at -20°C. The genomic DNA was used as template for PCR as described above with wild type and recombination specific primers from Tab. S 2. The products were run on agarose gel electrophoresis and expected bands for wild type and integration were analysed.
2.2.5.6 Isolation of clonal parasite line based on limiting dilution
To obtain an isogenic population of a new cell line, parasites get diluted to less than one parasite per dish in average, cultured and expanded. For this, the parasitemia of the used culture was precisely determined (count >1,000 erythrocytes) by a Giemsa smear. To determine the number of erythrocytes per/ml in the culture, 38 µl of RPMI 1649 complemented medium was mixed with 2 µl of the culture and gently resuspended. 10 µl were loaded on a haemocytometer and 5 of the inner 25 squares counted. The cell concentration was calculated by the following equation:
mean count ∗ number total squares ∗ dilution factor ∗ 10,000 = x RBC/ml
Afterwards, 1 x 106 cells were added to 10 ml of RPMI 1649 complemented medium and gently mixed. 50 µl (for 0.5 parasites/well) and 200 µl (for 2 parasites/well) were added to x ml of RPMI 1649 complemented medium with 2% haematocrit based on the following equation:
x ml = parasitemia of used culture x 10 ml
The clonal dilutions were incubated in the inner 60 wells of 96-well plates, while the outer wells were filled with 100 µl PBS. 100 µl of the previous diluted suspension were applied by a multichannel pipette to the 60 wells for the 0.5 parasite/well and 2 parasite/well dilution on separate 96-well plates. The suspension was all along kept suspended. Plates were closed and cultured in a humidified gas chamber. The gas
36 2ND CHAPTER - MATERIALS AND METHODS chamber was filled with fresh gas every 2-3 days. Every 6-7 days, 80 µl of medium was removed and the remaining erythrocytes resuspended in 90 µl of RPMI 1649 supplemented medium. Approx. 14 days after the start of the incubation, clones were expected. Slight colour change indicated a growing clonal culture. Some of the 0.5 parasites/well cultures were checked with Giemsa stains, and with a parasitemia >1% transferred to bigger culture volumes and scaled up and checked by diagnostic PCR. Parasite cultures with a parasitemia of >5% were cryo-preserved.
2.2.6 In vitro culturing of asexual P. falciparum 3D7 stages 2.2.6.1 Preparation of uninfected erythrocytes
Human blood was provided by the Red Cross. For blood with purified RBCs, the blood bag was split in smaller aliquots of 50 ml under sterile conditions. Scissors for the bag puncture need to be autoclaved and sterile conditions have to be applied as much as possible. The aliquots were stored at 4°C until the expiry date. For donations of whole blood, blood was centrifuged for 5 minutes at 1,500 rpm. The lowest layer of RBCs (underneath the buffy coat) was removed and transferred into fresh 50 ml tubes.
2.2.6.2 Serum deactivation
The serum was provided by the Red Cross and kept in plastic bags with a blood clot. The serum was aliquoted in 50 ml tubes without the clot and centrifuged at 2,000 rpm for 10 minutes. Based on the resulting supernatant the serum batch was assessed for use. Only serum with slightly transparent supernatant was used for culturing medium. Too cloudy or erythrocyte contaminated serum was avoided. Afterwards, the approved supernatant was transferred into new 50 ml tubes and heat-inactivated at 56°C for 1.5 hours. Stocks were kept at –20°C.
2.2.6.3 In vitro culture of P. falciparum
Asexual blood stage P. falciparum strain 3D7 parasites were cultured within human erythrocytes blood type 0+ in continuous culture using a method adapted from Maier and Rug [63]. The parasite culture was adjusted to 4% haematocrit in RMPI 1640 medium supplemented with 10 mM glucose solution, 480 µM hypoxanthine, 20 µg/ml gentamicin and serum from ALBUMAX II (0.375%, w/v) and heat inactivated
37 2ND CHAPTER - MATERIALS AND METHODS human serum (2,5%, w/v). Parasite culture was incubated in static petri dishes in gas chambers flooded with a culture specific gas composition of 94% nitrogen, 5% carbon dioxide and 1% oxygen and kept in at 37°C. Every 48-72h the parasitemia was estimated as below. The cells were split to 0.2% parasitemia with new adjusted 4% haematocrit in prewarmed culturing medium.
To synchronise the parasite culture to ring stages, the culture requires a minimum percentage of ~1% ring stages. This synchronisation was achieved by pelleting the culture for 4 minutes at 1,500 rpm, resuspension of 400 µl pellet with 2-5 ml prewarmed 5% sorbitol and incubation of the resuspension at 37°C for 10 minutes. After an additional pelleting for 5 minutes at 1,200 rpm, the synchronised pellet was resuspended in prewarmed RPMI medium with supplements. After 24 hours, the culture was monitored and split to 0.2%.
2.2.6.4 Assessment of parasitemia in an in vitro culture of P. falciparum
To monitor the development of the in vitro culture of P. falciparum the percentage of infected erythrocytes among all erythrocytes counted (parasitemia) was determined. 100-1,000 µl of the in vitro culture were briefly spun down in a reaction tube and a few µl of the pellet taken up with a pipette tip and transferred to a fresh glass slide. By using another glass slide the blood drop was evenly smeared over the glass slide with a slow and constant movement. After air drying, the slide was fixed and in 100% methanol for 10-30 seconds and stained with a 10% Giemsa solution for 5-10 minutes. After washing off the Giemsa solution under running water, the slide was dried and ready for microscopy. To count the parasitemia, parasites were observed under a bright-field microscope with 1,000-fold magnification using immersion oil directly on the stain. The overall condition of the culture was checked for contaminations, extracellular parasites, the condition of erythrocytes and parasites. Dependent on the necessary accuracy of the count, iRBC were counted per 500-1,000 total RBC and parasitemia was calculated in precent.
2.2.6.5 Thawing of P. falciparum stabilates
To start a new culture, cryo-stabilates were removed from liquid nitrogen and immediately thawed in a 37°C water bath. Afterwards, the stabilate was transferred
38 2ND CHAPTER - MATERIALS AND METHODS into a 50 ml tube and the volume was estimated. Dropwise, 12% NaCl in PBS (w/v) was slowly added under constant swirling into the tube. After an incubation of 5 minutes, 1.6% NaCl (w/v) was added drop-wise while constantly swirling the tube. Afterwards, 0.9% NaCl (w/v) with 0.2% glucose in PBS (w/v) were added again drop- wise while constantly swirling the tube. The resulting solution was centrifuged at 2,000 rpm for 5 minutes and the supernatant discarded. The pellet was resuspended in 10 ml RMPI 1640 without supplements while adding it very slowly into the first few millilitres. After an additional centrifugation at 2,000 rpm for 5 minutes the supernatant was discarded and the pellet resuspended in 10 ml RPMI 1649 with supplements. The suspension was transferred to a petri dish and cultured as described. After 24 hours, parasitemia was assessed as described above and adjusted to 1-2% with fresh RPMI 1649 with supplements and 4% haematocrit. After additional 48 hours, assessment of parasitemia and adjustment to 1-2% was repeated. When the culture remained stable, culturing methods were applied as described.
2.2.6.6 Saponin lysis
To isolate parasites by lysing the erythrocytes via a saponin lysis, the medium was resuspended and removed from a petri dish and centrifuged for 5 minutes at 1,500 rpm. The pellet was resuspended in 3 volumes of 0.15% saponin, which had been previously cooled on ice. The resuspension was transferred to a new tube and incubated on ice for 15 minutes. Every few minutes the suspension was inverted. After pelleting the parasites for 1 minute at 6,500 rpm, the supernatant was discarded and the pellet resuspended in 1 ml of ice-cold PBS. This step was repeated and again centrifuged for 1 minute at 6,500 rpm to get a parasite pellet of approx. 100 µl.
2.2.6.7 Cryopreservation of P. falciparum parasites lines
To cryo-preserve a stable culture, the overall condition of the culture was initially assessed by a Giemsa stain. It is recommended to freeze the parasites in a stable and well growing time point with a high parasitemia of ring stages (> 1%). The culture was spun down at 1,500 rpm for 5 minutes and the supernatant was removed. Two volumes of a sterilised freezing solution containing 28% Glycerol (v/v), 3% Sorbitol (v/v), 0.65% NaCl in tissue culture water was added drop-wise to one volume of the previous pellet. Meanwhile, the tube was swirled to keep the culture resuspended while adding the
39 2ND CHAPTER - MATERIALS AND METHODS freezing solution. The suspension was allowed to settle for 5 minutes and 0.5-1 ml were transferred to a freezing vial without creating bubbles nor wetting the rim of the tube. The tube was closed tight. To allow an even freezing, the tube was frozen in liquid nitrogen using forceps. Completely frozen tubes were kept and stored in liquid nitrogen.
2.2.7 P. falciparum SYBR safe-based in vitro growth assay
For an assessment of wild type-like asexual growth of the parasite, a SYBR safe based growth assay was established. The following procedures were performed on a wild type culture and the studied parasite line in parallel. To get a very synchronous ring stage culture, the cultures were sorbitol synchronised twice in an interval of 12-16 hours. Two days after the second synchronisation the cultures were monitored by a Giemsa stain to check for ring stages and a precisely determined parasitemia (count >1,000 erythrocytes). The cultures were centrifuged at 1,500 rpm for 5 minutes and the supernatant discarded. With fresh red blood cells, the culture was diluted to 1% parasitemia with 2% haematocrit with fresh erythrocytes and RMPI 1640 complemented medium. In a 96-well plate the outer wells were filled with PBS. On the same plate, 100 µl per well of the adjusted wild type or strain of interest were applied on 27 wells. For the remaining 6 wells, an uninfected 2% haematocrit culture was applied as a background control. The 96-well plates were incubated in a humidified gas chamber and flooded with the same gas composition used for normal culturing. After an incubation of 72 hours, the 96-well plates were frozen to stop the experiment. For the SYBR safe growth measurements, the plates were thawed and mixed with 100 µl of 1:5,000 SYBR safe dilution in lysis buffer. 100 µl of the gained suspension was transferred to a new 96-well plate and measured in a plate reader under 490nm excitation and 520nm emission with self-adjusted gain. The mean output of the erythrocyte only background was subtracted from the parasitised wells and fluorescence intensity of the wild type and studied parasite line was compared.
40 3RD CHAPTER – FT2
3rd Chapter: An apicoplast-resident folate transporter is essential for sporogony of malaria parasites 3.1 Introduction
In Plasmodium, folate serves as a vital co-factor which is required for one-carbon transfer reactions during DNA synthesis. Additionally, it is involved in the formation of methionine and used to initiate mRNA translation as well as protein and DNA methylation [64]. Accordingly, the importance of folate metabolism for Plasmodium parasites is evident from the past therapeutic exploitation of the biosynthetic enzymes of this pathway for the treatment of chloroquine-resistant malaria [65, 66]. The active ingredients of the antifolate combination drug Fansidar®, pyrimethamine and sulfadoxin, synergistically interfere with the parasite’s dihydrofolate reductase– thymidylate synthase (DHFR-TS) and dihydropteroate synthase (DHPS) enzymes, respectively, causing rapid parasite clearance upon treatment [67]. Despite the swift and global spread of antifolate resistance, Fansidar® is still used for intermittent preventive treatment, especially during pregnancy, which improves maternal and birth outcomes in malaria endemic areas [68]. The parasiticidal activity of antifolates also extends to gametocyte and liver stages, underscoring the universal essentiality of this co-factor for replicative and transmission stages in the mammalian host [69-73].
Much less is known about the parasite’s folate requirements during oocyst development and sporulation. A recent genetic investigation of folate synthesis and salvage during Plasmodium life cycle progression showed that the parasite’s folate requirements are dependent on both de novo synthesis and salvage from host diet [74]. The same study suggested that folate requirements during mosquito infection are rather modest compared to liver and blood stages, since the interruption of de novo synthesis and an excluded salvage from the hosts led to normal midgut infection and sporozoite development. Nonetheless, the increase in DNA content during sporogony is dramatic since a single oocyst forms several hundred new sporozoites after nuclear division [25, 75]. Therefore, a folate dependency during DNA synthesis in the Anopheles vector is very likely.
Even prior to their identification, folate transporters have already been discussed as lucrative drug targets [76]. As of yet, two folate transporters, called FT1 (e.g. PF3D7_0828600
41 3RD CHAPTER – FT2
and PBANKA_0702100), and FT2 (e.g. PF3D7_1116500 and PBANKA_0931500), were identified in the Plasmodium genome datasets. Both proteins contain two domains with high similarity to the Leishmania biopterin transporter 1 (BT1), a defining signature of BT1 transmembrane proteins (PFAM: PF03092) [77, 78], and belong to the Folate-Biopterin Transporter (FBT) Family, a member of the major facilitator superfamily (MFS) [41]. Via heterologous expression in Xenopus laevis oocytes and Escherichia coli bacteria by Salcedo-Sora and colleagues, FT1 and FT2 have shown to facilitate the transport of folates. However, the major prevalent folate, 5-methyltetrahydofolate, showed to be only transported by FT2. Other folates, and precursors like folic acid, folinic acid, pABA and pABAGn showed to be a substrate of FT1 and FT2, with pABA being the only cargo transported in physiological rates. Using polyclonal antisera of uncharacterized specificity, the P. falciparum orthologs of FT1 and FT2 were shown to localise to the parasite’s periphery and undefined internal membranes, implicating a folate and precursor salvage from the host in parallel to endogenous folate de novo synthesis [79].
For the folate synthesis, the parasite synthesises or imports the central precursor para-aminobenzoate (pABA) to fuse it with a pterin moiety derived from purine metabolism, followed by poly-L-glutamylation [64]. Evidence from other cell systems indicates that folate metabolism is highly compartmentalised, including reaction sequences in the cytosol, mitochondrion, plastid, vacuole and nucleus [80, 81]. Malaria parasites harbor two endosymbiotic organelles, the mitochondrion and a relic non-photosynthetic plastid, known as the apicoplast [82, 83]. Folate and/or intermediate metabolites are expected to enter these organelles; however, the identity of the involved transport proteins remains unknown.
P. berghei harbors FT1 and FT2 orthologues as well and enables access to a full life cycle approach to identify functional features beyond blood stages and to transfer the findings to the more relevant P. falciparum parasite. Using a combination of experimental genetics and quantitative live-cell microscopy, we investigate spatio- temporal expression of P. berghei FT2 and identify at which point during live cycle progression it fulfils a functional role.
42 3RD CHAPTER – FT2
3.2 Results 3.2.1 FT2 is broadly expressed throughout the life cycle and localises to the apicoplast For an initial assessment of FT2, the localisation and timing of expression of a fluorescently tagged FT2 protein were analysed. To this end, an isogenic transgenic P. berghei line expressing endogenous FT2 fused to a mCherry-3x-Myc tag was generated and selected (Fig. S 3 a). Successful generation of the FT2 locus was confirmed by diagnostic PCR (Fig. S 3 b top, cell-line produced by J. M. Matz). Microscopic examination of the fluorescently labelled FT2 during the P. b e r gh e i life cycle uncovered a striking organellar localisation, reminiscent of either the mitochondrion or apicoplast (Fig. S 3 c). The morphology of organelle branching is indicative of a more apicoplastidial structure than mitochondrial structure, and we tested for apicoplast targeting. Therefore, another independent line employing the same gene targeting strategy was generated, but this time parasites carried an additional apicoplast GFP-cassette for co-localisation by live fluorescence microscopy (Fig. S 3 a, cell-line produced by J. M. Matz). After confirmation of the correct integration (Fig. S 3 b bottom), spatio-temporal expression analysis of FT2-mCherry and the apicoplast GFP signals confirmed a distinct localisation of P. b er g h e i FT2 to the apicoplast (Fig. 3 a and b). FT2-mCherry was found to be expressed and overlapping with api-GFP signal during all parasite stages. However, in early oocysts this expression appears to be comparably low (Fig. 3 b).
Additionally, a further fraction of FT2-mCherry independent of the api-GFP signal in segmented schizonts was detected. The signal appears with comparable weaker fluorescence intensities at the apical prominence of merozoites (white arrowhead, Fig. 4 a). Similarly, sporozoites show an additional api-GFP independent fraction. Derived from the forward movement of the sporozoite, time lapse live imaging of motile sporozoites suggests the additional signal to localise again at the apical parasite pole (white arrowhead, Fig. 4 b; direction of movement indicated by white arrow). In late oocysts a non-plastidial signal can be seen as well, which most likely comes from the apical fraction of intra-oocystic sporozoites (white arrowhead oocysts d17, Fig. 3 b).
Together, this analysis uncovered an unanticipated localisation of P. b e r g he i FT2 to the apicoplast with a striking dual localisation at the apex of invasive merozoites and sporozoites. Furthermore, the expression during parasite life cycle progression appears to be continuous with potential differences in expression intensities.
43 3RD CHAPTER – FT2
DIC FT2::tag (a) FT2::tag api-GFP DAPI api-GFP
ring stage
trophozoite
schizont blood stages
segmenter
gametocyte
(b)
ookinete
oocysts d10 and EEFs
oocysts d17 mosquito stages
sporozoite
EEF 48h
44 3RD CHAPTER – FT2
Fig. 3 FT2 is an apicoplast protein and broadly expressed during Plasmodium life cycle progression.
(a, b) ft2-tagapiGFP parasites were imaged live at different stages. Shown are representative images of blood stages (a) and mosquito and EEF stages (b): a merge of differential interference contrast images (DIC) with Hoechst 33342 nuclear stain (DAPI, blue, 1st column), the fluorescent signals of tagged FT2 (greyscale, 2nd column), the apicoplast marker (api-GFP, green, 4th column) as well as a merge of tagged FT2 and api-GFP signals (red and green, 3rd column). Merge of tagged FT2 and api-GFP co-localises in all stages besides early oocysts. Note dual localisation of an additional FT2 fraction observed in non-apicoplast structures in segmented schizonts, sporulating oocysts and in sporozoites (white arrowheads). Extraparasitic red fluorescence within EEFs is due to autofluorescence of host cell vesicular compartments and was also observed in non-fluorescent wild type parasites and uninfected host cells (compare wild type EEFs Fig. S 4 ). Bars, 10 µm.
(a) DIC DIC api-GFP DIC FT2::tag DAPI FT2::tag FT2::tag
segmenter
(b)
0 seconds 36 seconds
FT2::tag
DIC
FT2::tag
Fig. 4 FT2 shows dual localisation to the apicoplast and to the apical end in segmented schizonts and sporozoites. (a) Shown are representative images of segmented schizonts: merge of DIC with Hoechst 33342 nuclear stain (DNA, blue, 1st column), DIC only (2nd column), merge of DIC with fluorescent signals of tagged FT2 (red, 3rd column), fluorescent signals of tagged FT2 in greyscale (4th column) and merge of apicoplast marker (api-GFP, green) with fluorescent signals of tagged FT2 (red, 4th column). Dual localisation overlaps apical 3D structure in DIC (white arrowhead) of segmented schizonts. (b) Shown is a sporozoite recorded during motility for a time-lapse of 36 seconds, including the signal of tagged FT2 (red, top) and a merge of DIC and tagged FT2 (greyscale and red, bottom). Arrow indicates movement during time-lapse. Bars, 10 µm.
45 3RD CHAPTER – FT2
3.2.2 Deletion of FT2: normal blood propagation but arrest of transmission by loss of sporulation. Since the expression of FT2 in the parasite’s life cycle is predominant, functional essentiality was investigated by the deletion of FT2 (Fig. 5 a). Using a replacement strategy based on double homologous recombination, P. b e rg h e i parasites lacking the FT2 gene were generated and isolated (performed by J. M. Matz). Additionally, the ft2– parasites express cytoplasmic GFP to allow identification by fluorescence microscopy and flow cytometry. An isogenic cell line was confirmed by diagnostic PCR with primers specific for recombination, highlighting the dispensability of FT2 for asexual proliferation in the mammalian blood stream (Fig. 5 b).
By examining quantifiable growth and development of the parasite, every stage in the life cycle was analysed to identify a potential developmental defect. In an intravital competition assay [60], mCherry expressing wild type parasites (Berred, P. berghei ANKA strain expressing cytoplasmic mCherry) and GFP-fluorescent ft2– were co-infected in the same mouse and analysed daily by flow cytometry. No short-term disadvantage in the rate of blood propagation was observed in individual animals (Fig. 5 c, performed by J. M. Matz). The conversion rate to sexual stages with FT2- deficiency appears unrestricted and shows no effect on developing fertile gametocytes (Fig. 5 d), quantities of exflagellation events of male gametocytes (Fig. 5 e), and ookinete conversion rates in ookinete cultures (Fig. 6 a). Together, these findings indicate that despite abundant expression, FT2 is dispensable for both sexual and asexual Plasmodium blood stage development.
Upon transmission to Anopheles stephensi mosquitoes, midgut colonisation showed to be unhindered for ft2– parasites (Fig. 6 b). Successful sporulation, the maturing of new sporozoites in oocysts, is an essential bottleneck in the parasite’s life cycle. By keeping midgut stages under surveillance for a period of 22 days after infection, dramatic interferences in the sporulation of ft2– oocysts were observed: for developing wild type oocysts, sporoblast-formation is observable between day 10 and day 20. The smooth and regular sporoblasts are formed after a detachment of plasma membrane from the capsule. During the formation, the cytosol becomes enclosed in the newly formed sporoblast, which is in wild type Bergreen parasites (P. b er g h e i ANKA strain expressing cytoplasmic GFP) represented by cytoplasmic expressed GFP (forming sporoblasts indicated by white arrowheads, Fig. 7 a top). Instead, FT2- deficient oocysts show an accumulation of extensive intraparasitic
46 3RD CHAPTER – FT2
(a) 5‘WT 3‘WT (b)
P1 P2 P3 P4 WT INT FT2 5‘ 3‘ 5‘ 3‘ - 2.0 kb ANKA P5 P6 - 0.5 kb GFP cassette drug cassette - 2.0 kb
ft2– - 0.5 kb 5‘INT 3‘INT
P1 P5 P6 P4 GFP cassette drug cassette
(c) (d) (e)
n.s. ) n.s. n.s. 1 10 20 10 100 d 15 8 10-1 6 10-2 10 4 10-3 5 Parasitemia (% ) 2 -4 WT 10 Exflagellations / fiel ft2– 0 0 – 3 4 5 6 7 WT ft2– WT ft2 Gametocyte conversion rate (% Days post infection
Fig. 5 Efficient blood propagation and sexual differentiation in the absence of FT2. (a) Replacement strategy for the deletion of P. berghei FT2. The locus was targeted with a replacement plasmid containing the 5′ and 3′ flanking regions (grey boxes), an expression cassette for cytoplasmic GFP fluorescence (green), and the drug-selectable hDHFR-yFcu cassette (blue). Shown are the wild type (WT) locus (top), the transfection vector (middle) and the recombined locus (bottom). WT-specific and INT-specific primer combinations are indicated by arrows. Molecular cloning and production of isogenic cell line performed by J. M. Matz. (b) Diagnostic PCRs of the WT locus (top), and of drug-selected and isolated ft2– parasites (bottom), using the primer combinations depicted in (a). Arrowheads show expected product size (details in Tab. S 1). (c, d) Efficient parasite proliferation in the murine bloodstream. (c) Intravital competition assay. Equal numbers of mCherry-fluorescent Berred WT and GFP-fluorescent ft2– blood stage parasites were co-injected intravenously into mice and peripheral blood was analysed daily by flow cytometry [60]. n.s., non-significant; two-way ANOVA to test for the two variables time and genotype; n=3. Performed by J. M. Matz. (d and e) Normal sexual differentiation in the absence of FT2. 107 WT or ft2– blood stage parasites were injected intravenously into mice and peripheral blood was analysed three days later for gametocyte conversion and (e) exflagellation events per microscopy field. Shown are the percentages of mature gametocytes among all blood stages, as identified in Giemsa-stained thin blood films and exflagellation centres per microscopic field. Shown are individual data points as well as mean values (lines). n.s., non-significant; Student’s t-test; n ≥ 6.
47 3RD CHAPTER – FT2
(a) n.s. (b) n.s.
100 1000
80 800
60 600
40 400
20 200 Ookinete conversion (% ) 0 Oocysts / infected midgu t 0 WT ft2– WT ft2–
d10 d17 (c) (d) n.s. * ) ) 3 3 150000 m m ( 20000 (
100000
10000 50000 Oocysts volume Oocysts volume 0 0 WT ft2– WT ft2–
Fig. 6 Ookinete conversion, mosquito midgut colonisation and oocyst development. (a) percentage of fully matured ookinetes among in vitro cultivated P28-positive parasites is normal in absence of FT2 (n=6), n.s., non-significant; Student’s t-test. (b) Normal mosquito midgut colonisation in the absence of FT2. Shown are oocyst numbers of individual infected midguts and the mean infection intensity (lines) on day 10 after the blood meal. n.s., non-significant; Student’s t-test; n ≥ 46 midguts from three independent blood meals. (c, d) Marked swelling of FT2-deficient mature oocysts. Oocysts were imaged live 10 (c) and 17 days after the blood meal (d). The area occupied by the oocyst in the micrographs was quantified and the oocyst volume was extrapolated assuming spherical proportions. n.s., non-significant; *, P<0.05; Student’s t-test; n ≥ 254 oocysts from 4 independent blood meals. Shown are individual data points as well as mean values (lines).
structures (yellow arrowhead, Fig. 7 a, bottom) which appear to result from the clumping of cellular components. It can be seen that these structures do not present the described typical sporoblast features, like a smooth surface or enclosure of cytoplasmic GFP. Differential interference contrast microscopy reveals that the displayed structures have rough surfaces, and are potentially made up of several smaller vesicles which increase in size by fusion. For late oocysts, complete vesiculation of the oocysts is observed (Fig. 7 b, d22 bottom). Cytoplasmic GFP
48 3RD CHAPTER – FT2 appears to be displaced by the structures (yellow arrowhead, Fig. 7 a, bottom). Additionally, live fluorescence imaging of the nuclei by Hoechst 33342 nuclear dye did not co-localise to the observed structures, while in wild type oocysts nuclei are inside of the sporoblast and aligned along the newly formed sporozoites (white and yellow arrowheads, Fig. 7 b WT top and ft2– middle). In addition to the described interference with the sporoblast formation, an inflated appearance of FT2-deficient parasites can be observed. In parallel to the normal increase in size during oocyst development, it can be shown that FT2-deficient oocysts become in average almost twice as voluminous as wild type oocysts by day 17 (Fig. 6 c and d). While the described aberrations (aggregate formation and lack of sporulation) can be seen in a small fraction of wild type oocysts as well (10.8% of day 17 oocysts, Fig. 7 c), FT2-deficient parasites show such alternations for the vast majority of oocysts in the late stages (58.0% of day 17 oocysts).
The defect results in only a very small proportion of all ft2– parasites to sporulate at all (2% compared to 50% in wild type, Fig. 7 d). Constantly, the tremendous diminished sporulation results in extremely low sporozoite quantities in the mosquito’s salivary glands (Fig. 7 e). Next, it was tested whether the few ft2–sporozoites produced are capable of eliciting patent blood infection and thus subjected naïve mice to the bites of infected mosquitoes or intravenous injection of 10,000 isolated sporozoites. In both approaches, the mice remained blood stage negative for a surveillance period of two weeks, while wild type re-infections exhibit blood stages three days post infection (Fig. 7 f). In addition, the few FT2-deficient sporozoites which are produced lost their ability for liver cell invasion: in an equal mix of 5,000 mCherry-fluorescent wild type and 5,000 GFP-fluorescent ft2– sporozoites, applied to cultured huh-7 cells, only mCherry-fluorescent wild type liver stages could be observed (Fig. 7 g).
49 3RD CHAPTER – FT2
Oocyst development (a)
DIC
WT
GFP
DIC
–
ft2
GFP
DIC DIC (c) (b) GFP n.s. *** GFP DAPI 60
40
WT
d 17 20 Abnormal oocysts (% ) 0 WT ft2– WT ft2– (d) d10 d17
d 17 60 ***
–
ft2 40
d 22 20
Sporulating oocysts (% ) 0 (e) (f) – WT ft2 ) (g) ) 15000 *** 1.0 o 100
10000 ft2– by bite 0.5 ft2– i.v. 50 5000 WT by bite
WT i.v. ft2– wilde type Sporozoites / mosquit 0 Parasite-free animals (% 0.0 0 WT FT2- 0 3 6 9 12 15 Days post infection Liver stages (% of all parasites expectedobserved
50 3RD CHAPTER – FT2
Fig. 7 Prominent swelling and cytoplasmic malformations in FT2-deficient oocysts. (a) Depicted are differential interference contrast (DIC) images of developing oocysts (day 10 till day 17, arrow indicates development), showing various stages of either sporulation (WT, top) or malformations in form of cytoplasmic accumulations (ft2–, bottom; DIC 1st and 3rd row and cytoplasmic GFP 2nd and 4th row). White arrowheads, sporoblast; yellow arrowheads, rough patches. (b) Intracellular malformations displace cytoplasmic GFP and nuclei in FT2-deficient oocysts. Shown are images of a sporulating WT oocysts at day 17 (top) and ft2– oocysts at day 17 and 22 after the blood meal (bottom), including a merge of the fluorescent signal of DIC and cytoplasmic GFP (green, 1st column), fluorescent signal of cytoplasmic GFP only (green, 2nd column), and a merge of DIC and Hoechst 33342 nuclear stain (DAPI, blue, 3rd column). Note the presence of cytoplasmic GFP within the sporoblast (white arrowhead) and the absence thereof from the rough patches (yellow arrowhead). Also, note the displacement of nuclei by the rough patches. At day 22 the ft2– oocysts are completely vesiculated. Bars, 10 µm. (c) FT2-deficient oocysts form large intracellular malformations. Shown is the percentage of abnormal oocysts containing intraparasitic accumulations like in (a) 10 and 17 days after the blood meal. n.s., non- significant; ***, P<0.001; Student’s t-test; n=4 independent blood meals. (d) Oocyst sporulation is severely impaired upon loss of FT2. Sporulating oocysts were quantified on day 17 after the blood meal. ***, P<0.001; Student’s t-test; n=4 independent blood meals. Percentages indicate the mean sporulation frequency. (e) Quantification of salivary gland-associated sporozoites 21 days after the blood meal. Depicted are individual averaged sporozoite burdens from ~50 mosquitoes per feeding (dots) as well as mean values. ***, P<0.001; Student’s t-test; n=4 independent blood meals. (f) Sporozoites originating from FT2-deficient oocysts do not elicit malaria. Kaplan-Meier analysis of time to development of patent blood infection. Naïve mice were intravenously injected with 10,000 WT or ft2– sporozoites, or were subjected to bites by mosquitoes that had received blood meals from mice infected with WT or ft2– parasites. Peripheral blood was monitored daily by microscopic analysis of Giemsa-stained thin blood films; n=3 mice from two independent feeding experiments. (g) Sporozoites originating from FT2-deficient oocysts are non-invasive. Huh-7 cells were inoculated with an equal mix of 5,000 mCherry-fluorescent Berred WT and 5,000 GFP-fluorescent ft2– sporozoites obtained from independent preparations. Shown are expected (left) and observed ratios of WT and ft2– liver stage parasites (right), as quantified by live fluorescence microscopy 48 hours after inoculation. n=4 wells with 30,000 Huh-7 respectively from one infection experiment.
51 3RD CHAPTER – FT2
3.2.3 Rescue of defect by ectopic expression of FT2::mCherry in SIL6 in FT2- deficient and wild type parasites. To exclude indirect effects due to secondary site mutations, FT2-deficient parasites were complemented with FT2 by transfecting a FT2-mCherry-3xMyc fusion protein expression cassette under the CPN20 promoter in the SIL6 locus into a FT2- deficient parasite line (ft2comp, Fig. 8 a, cell line generated by J. M. Matz). Additionally, overexpression of FT2 was induced by introducing the same transfection approach in wild type parasites harbouring endogenous FT2 (ft2over, cell line generated by J. M. Matz). The correct integration and isogenic cell lines were approved by diagnostic PCR (Fig. 8 b). Live fluorescence microscopy showed a re-localisation of complemented FT2::mCherry (Fig. 8 d) with comparable features like in previous localisation- experiments (Fig. 3 a). Furthermore, the two cell lines were assessed for developmental defects, like observed in the FT2-deficient cell lines. The sexual development showed normal progression (Fig. 9 a and b) and mosquito midgut colonisation occurred in wild type comparable quantities (Fig. 9 c). Size increases and inflation of oocysts were restored in day 17 oocysts and showed a trend for smaller volumes in day 17 ft2comp oocysts (Fig. 9 d and e). For both cell lines sporoblast formation showed wild type-like entities (white arrowheads, Fig. 8 c and e) which lead to the formation of sporozoite-rich oocysts and sporozoites extractable from salivary glands (Fig. 8 c 3rd and 4th row, e 2nd and 3rd row). The quantification of sporulating oocysts, sporozoites extracted from salivary glands and the quantification of FT2- deficiency associated abnormalities in oocysts confirmed a restoration of the defect from ft2– parasites (Fig. 9 f, g and h). Finally, the infection of naïve mice with 30,000 sporozoites isolated from salivary glands of WT and ft2comp-infected mosquitoes showed blood stage parasites in the peripheral blood in parallel to wild type development (Fig. 9 i).
In summary, the defect and life cycle arrest induced by the deletion of FT2 was entirely restored by ectopic expression of FT2 in SIL6, and overexpression by a second copy did not lead to secondary effects.
52 3RD CHAPTER – FT2
WT SIL6 FT2 (a) (b) INT WT INT P1 P2 SIL6 WT 5‘ 3‘ 5‘ 3‘ 5‘ 3‘ PBANKA_061210 PBANKA_061220 - 2.0 kb ANKA
5‘CPN20 FT2 tag drug cassette - 0.5 kb - 2.0 kb FT2 5‘INT 3‘INT ft2over
P1 P3 P4 P2 - 0.5 kb 5‘CPN20 FT2 tag drug cassette - 2.0 kb
ft2comp
- 0.5 kb
ft2 (c) ft2 comp comp (d) DIC DIC DIC FT2::tag GFP FT2::tag GFP DAPI GFP DIC, DAPI
d10
trophozoite
(e) ft2over DIC DIC GFP d17 GFP DAPI
oocysts
d10 oocyst
d17
d17 oocyst
sporozoite
sporozoite
Fig. 8 Ectopic expression in ft2– line to rescue sporogonic defects and to assess overexpression of FT2. (a) Recombination strategy to complement ft2– parasites with FT2::tag or to overexpress FT2::tag in wild type parasites. Negative selection of ft2– parasites (from Fig. 5 a) with 5-fluorocytosine removes large parts of the drug-selectable cassette through homologous recombination of a duplicated sequence. The yielded pyrimethamine-sensitive ft2– parasites are used for the targeting of the silent locus SIL6 with a plasmid harbouring the CPN20 promoter driving expression of mCherry-3xMyc-tagged (tag, red) P. berghei FT2. Additionally, the construct carries a drug cassette for positive selection with pyrimethamine (blue). Wild type locus (top), vector for transfection (middle) and recombined locus (bottom). Primer combinations for diagnostic PCR are indicated by arrows. (b) Diagnostic PCR for SIL6 with primer combinations from (a) and endogenous FT2 locus for wild type parasites, wild type parasites expressing FT2 in SIL6 (ft2over)
53 3RD CHAPTER – FT2
– and ectopic expression in ft2 line (ft2comp). Arrowhead indicate expected PCR products (details in – Tab. S 1) (c) Rescue of sporogonic defects of ft2 in ft2comp parasites. Shown are representative images of a merge of differential interference contrast (DIC) and cytoplasmic GFP (green, 1st column), cytoplasmic GFP only (green, 2nd column), and a merge of DIC and Hoechst 33342 nuclear stain (DAPI, blue, 3rd column) for ft2comp oocysts on day 10 and 17, as well as sporozoites isolated from salivary glands on day 23. Oocysts show wild type like sporoblasts (white arrowhead) and sporozoite-rich late oocysts (3rd row). Bar, 10 µm. (d) Localisation of ectopically expressed FT2. Shown is a live fluorescence micrograph of a representative ft2comp trophozoite, including a merge of DIC and cytoplasmic GFP (green, left), a merge of DIC, the fluorescent signal of tagged FT2 (red) and Hoechst 33342 nuclear stain (DAPI, blue, centre) and the fluorescent signal of tagged FT2 (greyscale, right). Bar, 5 µm. (e) No developmental impact on sporogony following FT2 – overexpression in ft2 parasites (ft2over). Shown are representative images of a merge of DIC and cytoplasmic GFP (green, 1st column), cytoplasmic GFP only (green, 2nd column), and a merge of DIC and Hoechst 33342 nuclear stain (DAPI, blue, 3rd column) for ft2over oocysts on day 10 and 17, as well as sporozoites isolated from salivary glands on day 23. Oocysts show wild type like sporogony and sporozoites. Bars, 10 µm.
54 3RD CHAPTER – FT2
(a) (b) (c)
50 100 1000
40 80 800
30 60 600
20 40 400
10 20 200 Ookinete conversion (% ) 0 0 Oocysts / infected midgu t 0 WT ft2comp ft2over WT ft2comp ft2over WT ft2comp ft2over Gametocyte conversion rate (% )
(d) (e) (f)
80 10000 30000 ) ) 3 3 m 8000 m 60 20000 6000 40 4000 10000 20 2000 d10 oocysts size ( d17 oocysts size ( 0 0 0 WT ft2comp ft2over WT ft2comp ft2over Sporulating oocysts d17 (% ) WT ft2comp ft2over (g) (h) (i) n.s. ) 80 15000 100 o WT ) 80 60 ft2comp 10000 60 40 40 5000 20 20 Sporozoites / mosquit Parasite-free animals (% Abnormal oocysts (% 0 0 0 WT ft2comp ft2over WT ft2comp ft2over WT ft2comp ft2over 0 5 d10 d17 Days
Fig. 9 Phenotyping of ectopic expression of FT2 in ft2– line to rescue sporogonic defects and to assess FT2 overexpression.
7 (a) 10 wild type, ft2comp and ft2over blood stage parasites were injected intravenously into mice and peripheral blood was analysed three days later for gametocyte conversion (a) and ookinete conversion (b). After natural transmission to the mosquito, normal midgut colonisation is observed (c). Swelling like in ft2– parasites cannot be observed for day 10 (d) and day 17 oocysts (e) but ft2comp parasites show a trend for smaller oocysts volumes at day 17. The area occupied by the oocyst in the micrographs was quantified and the oocyst volume was extrapolated assuming spherical proportions. Quantification of sporulating oocysts shows no impairment on oocyst’s sporulation. Line represents mean (f) and the presence of abnormal formations like in ft2– parasites are comparable to the trends in wild type parasites (g). Sporozoites can be isolated from mosquito salivary glands (h). n=1. (i) Kaplan-Meier analysis of time to development of patent blood infection. Naïve mice were intravenously injected with 30,000 sporozoites isolated from salivary glands of
55 3RD CHAPTER – FT2
WT and ft2comp-infected mosquitoes and peripheral blood was monitored daily by Giemsa blood smears. n.s., non-significant; Mantel-Cox test; n=9 mice from 3 independent feeding experiments.
3.2.4 Heterozygous oocysts after cross-fertilisation of ft2– and wild type parasites overcome defect and produce haploid ft2– sporozoites To address a potential impact of FT2 on the liver stage development of the parasite, FT2-deficient sporozoites were generated by performing a red-fluorescent wild type (Berred) and green-fluorescent ft2– parasite cross-fertilisation. Thereby, after sexual recombination in the mosquito midgut, heterozygous double-fluorescent WT x ft2– oocysts are formed which generate a mix of WT and ft2– sporozoites after different allele combinations for FT2/ft2– with the WT’s mCherry/SIL6 locus (Fig. 10 a). After liver-cell infection, the genotype can be concluded from the respective fluorescence by live fluorescence microscopy of infected Huh7-cells or - after infection of naïve mice - by flow cytometry of blood stages (ft2– parasites in green and green + red, WT parasites in red and without fluorescence, Fig. 10 a). Therefore, parallel infection of cultured Huh7 liver cells with a mixed sporozoite population resulted in the maturation of both WT and FT2-deficient liver stages. No differences in size for ft2– liver stage parasites can be observed (Fig. 10 b). Additionally, naïve mice were intravenously injected with the same mix of sporozoites. The emergence of WT and FT2-deficient blood stages, both three days following injection, was measured by flow cytometry of peripheral blood (Fig. 10 c and d). Therefore, no developmental delay or arrest was measured in liver stage development.
Together, these findings indicate that the life cycle arrest in the absence of FT2 is not caused by defective maturation during the liver stage development. It is rather rooted in the inefficient production of non-invasive sporozoites within the mosquito vector, underscoring an essential function of the apicoplast transport protein FT2 in Plasmodium sporogony.
56 3RD CHAPTER – FT2
(a) Oocysts Sporozoites Liver stages Blood stages genotype colour genotype colour colour colour (dipolid) (expressed) (haploid) (carry-over) (expressed) (expressed)
Chr. 6 - Chr. 9 ft2– GFP FT2-deficient Chr. 6 mCherry
WT mCherry Chr. 9 FT2 Chr. 6 Chr. 9 ft2– GFP FT2-deficient X Chr. 6 - Chr. 6 - – ft2 Chr. 9 FT2 expression Chr. 9 ft2– GFP FT2
Chr. 6 mCherry FT2 expression Chr. 9 FT2
Chr. 6 mCherry WT Chr. 9 FT2 Chr. 6 mCherry FT2 expression X Chr. 9 FT2 Chr. 6 mCherry WT Chr. 9 FT2
Chr. 6 - ft2– Chr. 9 ft2– GFP Chr. 6 - X arrest of – X Chr. 9 ft2 GFP transmission Chr. 6 - ft2– Chr. 9 ft2– GFP
(b) (c) (d) 24h 48h
) n.s.
1.0 ft2– WT
WT
2 2 48 µm 410 µm 0.5
–
ft2
Parasite-free animals (% 0.0 0 1 2 3 4 5 51 µm2 446 µm2 Days post infection
Fig. 10 Phenotypic rescue during mosquito infection restores ft2– sporozoite infectivity and reveals redundant functions of FT2 during liver stage development. Mosquitoes were fed on mice co-infected with mCherry-fluorescent Berred WT and GFP- fluorescent ft2– parasites, leading to cross-fertilisation and phenotypic rescue during mosquito infection. (a) Interchromosomal recombination during the mosquito stage. Genotypes and fluorescence properties of heterozygous Berred WT x ft2– oocysts (orange, top) homozygous Berred WT (red, middle), and homozygous ft2– oocysts (green, bottom), and of the resulting sporozoites, liver stages and blood stages. Depicted are the FT2 locus on chromosome (Chr.) 9 and the silent intergenic locus on chromosome 6. Berred WT parasites harbor the intact FT2 gene and express mCherry (mCh) from Chr. 6. ft2– parasites express GFP from the disrupted FT2 locus, while Chr. 6 is unaltered. Interchromosomal recombination of heterozygous Berred WT x ft2– during mosquito stage development yields four distinct genotypes for the haploid sporozoites emerging from heterozygous oocysts. Note that these sporozoites remain double-fluorescent due to protein carry-over from the oocyst. Sporozoites from homozygous ft2– oocysts are non-infective. For the quantitative analysis of mosquito-to-mouse transition, only fluorescent parasites were quantified. Thus, non-fluorescent liver- and blood stages were not detected. (b) Representative images of in
57 3RD CHAPTER – FT2 vitro cultivated WT and ft2– liver stages at different times after infection with sporozoites isolated from WT x ft2–-infected mosquitoes. Shown is a merge of cytoplasmic fluorescence (red, WT; green, ft2–), Hoechst 33342 nuclear stain (blue) and differential interference contrast images. Values represent the mean area occupied by liver stages in fluorescence micrographs. Bars, 10 µm. (c) Kaplan-Meier analysis of time to development of patent blood infection. Naïve mice were intravenously injected with 30,000 sporozoites isolated from salivary glands of WT x ft2–-infected mosquitoes and peripheral blood was monitored daily by flow cytometry. n.s., non-significant; Mantel-Cox test; n=9 mice from 3 independent feeding experiments. (d) Representative dot plot obtained by flow cytometry on day 3 after transmission. Double fluorescent parasites are due to interchromosomal recombination events during mosquito infection and denote FT2-deficient parasites.
58 3RD CHAPTER – FT2
3.2.5 FT2 ortholog in P. falciparum Since FT2 shows relatively high conservation between P. b er g h e i and P. falciparum (86% amino acid identity between FT2 orthologs), a potentially conserved function was analysed by re-assessing FT2’s localisation and an essential function in blood stages via experimental genetics on P. falciparum 3D7.
Disruption and partial deletion of FT2 have been approached using CRISPR/Cas9-mediated double-strand breaks with a pDC2 plasmid (comparable to Fig. 15 a), as well as conventional double homologous recombination with a pCC1 plasmid (Fig. 11 b and Fig. S 6 b). For CRISPR/Cas9-mediated genome editing two constructs with different gRNAs have been designed to induce a double-strand break at position 3293 and 3485 in FT2. The selection of specific gRNAs was met on optimised on-target and off-target scores identified after Hsu et al. [84] and Doench et al. [85] (Fig. 11 a). gRNA was cloned in the vector at the BbsI site identical as described for Pf3D7_1468600 below (Fig. S 2). The assembly of EcoRI and ApaI digested plasmid, drug cassette and the PCR produced homologous regions was performed via a one-step Gibson assembly. Sequencing primers annealing within reach of the fused fragments (compare Fig. S 2) have been used to check for correct assembly by Sanger sequencing. Transfections with different Pf3D7 wild type cell populations have been performed with each construct, cultured and monitored for a period of 60 days under drug pressure. No parasite culture was established after three transfections with the two gRNA designs respectively (Fig. 11 a, 1st and 2nd column).
The transfected pCC1 constructs designed for partial deletion of FT2 via double homologous recombination can be found in Fig. S 6 a. After the digest of the plasmid with SacII, EcoRI, SpeI and AvrII the homologous regions of P. falciparum FT2 PCR products were reassembled with the drug cassette and plasmid. After confirmation of the right assembly by Sanger sequencing with respective sequencing primers (purple, Fig. S 6 a), the plasmids were prepared for transfection and transfected three times with different Pf3D7 wild type parasite cultures and monitored under drug pressure for at least 60 days (recombination strategy Fig. 11 b). In two out of three cultures, the first parasites were observed after 24 and 32 days (Fig. 11 a, 3rd column). The established cultures underwent drug-cycling for positive and negative selection. Extracted DNA from parental parasite populations was used for diagnostic PCR with primer combinations indicated in Fig. 11 b. The parental lines showed the presence of wild
59 3RD CHAPTER – FT2
(a) CRISPR-Ca9 mediated disruption of P. falciparum FT2
position 3293 position 3485 Deletion of FT2 Tagging of FT2 MTP HR HR MTP HR HR by pCC1 vector by pGlux6 off-target score1: off-target score1: double- 50.0 50.0 episomal GFP remarks homologous on-target score2: on-target score2: tag of FT2 recombination 50.9 61.0
transfected 3 transfections 3 transfections in 2/3 transfections 3 transfections population unsuccessful unsuccessful after 24 and 32 unsuccessful after days (b) (c) 5‘WT 3‘WT
P1 P2 P3 P4 WT INT
MTP 5‘ 3‘ 5‘ 3‘ - 3.0 kb
Pf3D7 FT2 - 1.0 kb deleted P5 P6 - 0.5 kb drug cassette
WT INT AmpR CD cassette 5‘ 3‘ 5‘ 3‘ - 3.0 kb
5‘INT 3‘INT Pf3D7 - 1.0 kb P1 P5 P6 P4 - 0.5 kb drug cassette
Fig. 11 Experimental genetics approach to interrupt, delete or GFP-tag P. falciparum FT2. (a) transfection overview of deletion and FT2-tag strategies. CRISPR/Cas9-mediated interruption of FT2 with two different gRNA positions on P. falciparum FT2 (position 3293 and 3485) between the homologous regions (HR). Off-target and on-target calculation after 1: Hsu [84] et al. and 2: Doench et al. [85]. Both gRNA designs showed no cell population after 60 days of culturing in three transfection approaches respectively. Deletion of P. falciparum FT2 by double homologous recombination with a pCC1 vector led in two out of three transfections to cell populations. Episomal tagging of P. falciparum FT2 with GFP by a pGlux6 vector led in none of three transfections to an establishment of a population. (b) Recombination strategy for deletion of P. falciparum FT2 by double homologous recombination with the pCC1 vector. Shown are the wild type (WT) locus (top), the transfection vector (middle) and the recombined locus (bottom). The targeting plasmid carries an Ampicillin resistance cassette for molecular cloning (AmpR, green) and a drug cassette for positive selection (hDHFR, blue) flanked by homologous regions (grey boxes). Double homologous recombination design leads to a partial deletion of the gene. An additional CD cassette (yellow) allows negative selection for integration of the backbone of the plasmid. Primer combinations (P1- P6) on wild type locus allow diagnostic PCR to test for recombination and isogenic populations after drug-cycling. (c) diagnostic PCR to test for wild type (WT) or integration specific (INT) PCR products indicated in (b). pCC1 transfected cell line show only 3’INT integration and presence of wild type locus PCR products (top). This pattern was observed for both populations after drug cycling including negative selection of the transfectant population. Wild type parasites show 5’WT and 3’WT bands and no integration specific products (bottom). Arrowheads indicate expected product size (details in Tab. S 2).
60 3RD CHAPTER – FT2 type specific PCR products (5’WT and 3’WT) and correct integration for the homologous region at the 3’ end (3’ INT). At the 5’ homologous region (5’INT) no PCR product was amplifiable (Fig. 11 c, top). Therefore, a deletion of the targeted gene fragment was unsuccessful. This pattern was observed for both of the transfected cell lines.
For a re-evaluation of P. falciparum FT2’s localisation, FT2 was episomal expressed under the CRT promoter fused to a GFP tag on a pGlux6 plasmid (Fig. S 6 b). Since the PCR amplification of P. fa lciparum FT2 under its endogenous promotor was not possible, only the gene was amplified from genomic DNA. After the digest with XhoI and KpnI the PCR product was ligated in the XhoI and KpnI digested pGlux6 plasmid. After confirmation of the correct ligation by Sanger sequencing with the sequencing primer indicated in Fig. S 6 b (purple) the vector was prepared for transfection. Three transfections with different Pf3D7 wild type parasites were performed. The cultures were monitored under puromycin drug pressure for a period of 60 days. No cell population was established in any of the transfections.
61 3RD CHAPTER – FT2
3.2.6 Methionyl-tRNA formyltransferase (FMT): a folate dependent pathway in the apicoplast To assess if the observed defect in ft2– parasites during sporogony is driven by a lack of folates within the apicoplast, Plasmodium pathways which potentially require the presence of plastidial folate were approached. The relict plastid of photosynthetic ancestry still harbours bacterial-type metabolic pathways and housekeeping pathways [6]. Therefore, a methionyl-tRNA formyltransferase (FMT) of bacterial origin, which initiates translation by charging a specialised tRNA with formylmethionine, has been shown to localise in Toxoplasma in the apicoplast [86]. Since in Plasmodium FMT performs the formylation with a formyl functional group originating from 10- formyltetrahydrofolate [64], impairments in the plastid’s folate household could lead to the observed defects. In reverse, if the defect is based on a reduced or absent formylation of methionyl-tRNA, the genetic deletion of FMT should potentially reproduce the defect of FT2-deficient parasites. Transgenic FMT-deficient parasites and parasites with endogenous FMT fused to mCherry-3xMyc with simultaneous expression of GFP in the apicoplast, were generated by J. M. Matz following a design identical to Fig. S 3 a and Fig. 5 a.
Initially, the spatio-temporal FMT::mCherry fusion protein expression is assessed for the entire P. be r g he i life cycle progression. Strikingly, the FMT::mCherry signal (white arrowhead, Fig. 12 c) co-localises with the api-GFP signal (yellow arrowhead, Fig. 12 c) and shows abundant expression in the EEFs apicoplast. Remarkably, this expression appears to switch off during blood stage infections and remains below the detectable threshold in ookinetes, as well as oocysts and sporozoites (Fig. 12 a and b). A lack of expression implies no functional role during the respective stages. Nevertheless, phenotyping of the entire life cycle progression was performed (Fig. 13 a-h and Fig. S 7 a-e). fmt– parasites showed normal blood stage propagation (Fig. S 7 a, performed by J. M. Matz) as well as normal gametocyte conversion (Fig. S 7 b) and normal ookinete formation (Fig. S 7 c) compared to wild type parasite growth. The colonisation of developing midgut oocysts was counted and showed no major differences (Fig. S 7 d). Day 10 and day 17 oocysts have been imaged and the area occupied by cytoplasmic mCherry was measured (Fig. 13 e and f). The area was transformed to volume assuming spherical proportions of the oocysts. The inflation of the volume at day 17 like in ft2– parasites was not reproduced (Fig. 13 f). When oocyst sizes are compared on the area level instead of a volume
62 3RD CHAPTER – FT2
DIC FMT::tag (a) api-GFP FMT::tag DAPI api-GFP
ring stage
trophozoite
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segmenter
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(b)
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63 3RD CHAPTER – FT2
(c) DIC FMT::tag api-GFP FMT::tag DAPI api-GFP
EEF 48h EEFs
EEF 66h
Fig. 12 FMT is an apicoplast protein and exclusively expressed in Plasmodium EEFs.
(a, b, c) Expression of FMT throughout the P. berghei life cycle. fmt-tagapiGFP parasites were imaged live at different stages. Shown are representative images of blood stages (a), mosquito stages (b) and EEF stages (c): a merge of differential interference contrast images (DIC) with Hoechst 33342 nuclear stain (DAPI, blue, 1st column), a merge of the mCherry fluorescent signals of tagged FMT with the apicoplast marker api-GFP (mCherry: red, api-GFP: green, 2nd column), the apicoplast marker api-GFP (green, 3rd column) as well as the mCherry fluorescent signals of tagged FMT only (red, 4th column). Only in EEFs FMT::mCherry shows an expression (white arrowhead) and co-localisation (yellow arrowhead) with the apicoplast GFP signal. Bars, 10 µm. transformation, a similar outcome is shown (Fig. S 7 e). Next, the morphological classification of sporoblast formation or vesiculation is based on features as described for ft2– parasites. Early-stage oocysts on day 10 do not show such differentiations and were therefore excluded from the quantification (Fig. 13 a, d10 1st row). Sporoblasts, which enclose cytoplasmic mCherry and form sporozoites (Fig. 13 a, d17, 2nd row, white arrowhead: sporoblast enclosing cytoplasmic mCherry, red arrowhead: DNA of sporozoites radial surrounding sporoblast), as well as abnormal oocysts with rough- surfaced vesicles displacing cytoplasm and DNA (Fig. 13 a, d17 3rd row, white arrowhead: displacement of cytoplasmic mCherry, red arrowhead: displacement of DNA) are observed in live fluorescence microscopy of FMT-deficient d17 oocysts. Furthermore, sporulation (Fig. 13 b) and the appearance of ft2– like abnormalities (Fig. 13 c) occur in comparable intensities like in wild type parasites. Extraction of sporozoites from the mosquito’s salivary glands on day 22 after blood meal generates sporozoites in wild type-like figures (Fig. 13 d). Finally, in reinfection experiments it was tested whether the fmt– sporozoites are capable of eliciting patent blood infection and
64 3RD CHAPTER – FT2
(a) DIC DIC (b) (c) mCherry mCherry DAPI
80 80
60 60
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– 20 20
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0 Abnormal oocysts d17 (% ) 0 Sporulating oocysts d17 (% ) WT fmt– WT fmt–
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(d) (e) (f)
15000 10000 30000 ) ) 3 3 m 8000 m 10000 20000 6000
4000 5000 10000 2000 d10 oocysts size ( d17 oocysts size ( Sporozoites / mosquit o 0 0 0 WT fmt– WT fmt– WT fmt–
(g) (h) ) n.s. 100 WT ) 80 fmt– 3 m 4000 60
40 2000 20 EEF volume (
Parasite-free animals (% 0 0 0 5 10 fmt– WT fmt– WT fmt– WT Days 24h 48h 69h
Fig. 13 Loss of FMT does not lead to a copy of the defect observed in FT2-deficient parasites. (a) Shown are images of fmt–oocysts at day 10 (top) and day 17 after the blood meal (bottom), including a merge of the fluorescent signal of cytoplasmic mCherry and DIC (1st column), fluorescent signal of cytoplasmic mCherry only (greyscale, 2nd column), and a merge of DIC and Hoechs t 33342 nuclear stain (DAPI, blue, 3rd column). Both sporulation and abnormal oocyst formation are observed in fmt– oocysts on day 17 after blood meal. The rough patches show comparable entities to ft2– oocysts like displacement of cytoplasmic mCherry and nuclei (yellow arrowhead: displacement of cytoplasm, red arrowhead: displacement of DNA). Sporoblast formation occurs without any morphological damage and encloses cytoplasm (white arrowhead) and forms new sporozoites aligning around the sporoblast (green arrowhead: DNA of sporozoites). Bars, 10 µm. (b) Oocyst sporulation is not severely impaired upon loss of FMT. Sporulating oocysts were quantified on day 17 after the blood meal. Percentages indicate the mean sporulation
65 3RD CHAPTER – FT2 frequency; n=1. (c) Shown is the percentage of abnormal oocysts containing intraparasitic accumulations like in (a) 17 days after the blood meal; n=1. (d) Quantification of salivary gland- associated sporozoites 21 days after the blood meal. Depicted are individual averaged sporozoite burdens from ~50 mosquitoes per feeding (dots). fmt–oocysts do not show phenocopy of ft2– oocysts, since sporozoites are being produced; n=1. (e, f) Oocysts size was measured live 10 (e) and 17 days after the blood meal (f). The area occupied by the oocyst in the micrographs was quantified and the oocyst volume was extrapolated assuming spherical proportions from 1 blood meal. Shown are individual data points as well as mean values (line). Swelling in late stages does not occur. (g) Kaplan-Meier analysis of time to development of patent blood infection. Naïve mice were intravenously injected with 10,000 sporozoites isolated from salivary glands of WT and fmt–- infected mosquitoes and peripheral blood was monitored daily by flow cytometry. n.s., non- significant; Mantel-Cox test; n=9 mice from 3 independent feeding experiments. (h) In vitro cultivated WT and fmt– liver stages at different times after infection with sporozoites isolated from WT and fmt–-infected mosquitoes. EEFs were imaged live 24h, 48h and 69h after Huh7 infection with sporozoites. The area occupied by the EEF in the micrographs was quantified and the oocyst volume was extrapolated assuming spherical proportions from sporozoites of one blood meal. Minor size differences are observed for late fmt– liver stages. Shown are individual data points as well as mean values (line).
thus subjected naïve mice to intravenous injection of isolated sporozoites. The mice showed fmt– blood stages at parallel time points to wild type sporozoite infections (Fig. 13 h). Additionally, Huh7 liver cells were infected with extracted sporozoites to test for invasion and to trace the EEF development. 24, 48 and 69 hours after infection live imaging of infected Huh7 cells allowed size measurements and showed the enlargement of the parasites. Minor reduction in the size of fmt– EEFs can be observed (Fig. 13 g). Note, that statistics was not performed, since we tested only one biological sample to identify successful sporogony as a qualitative output in order to identify a copy of the phenotype observed in FT2-deficient parasites.
In summary, the data confirm that Plasmodium FMT localises in the apicoplast as it was observed in Toxoplasma. Additionally, we detected an exclusive expression in liver stages only. Deletion of FMT demonstrated no essential role during the entire life cycle progression, including liver stages. Beyond that, growth defects upon deletion of FMT as observed in ft2– parasites were not reproduced.
66 3RD CHAPTER – FT2
3.3 Discussion Here, we demonstrated that FT2 is a transporter of the apicoplast with essential functions during Plasmodium oocyst sporulation. A previous study claimed localisation of the P. falciparum FT2 orthologue to the parasite’s periphery, based on immunofluorescence analysis with polyclonal anti-peptide antisera [79]. By contrast, genetic evidence for an apicoplast localisation of FT2 in P. b er g h e i was provided. Heterologous expression of tagged FT2 restored efficient sporulation and sporozoite infectivity in parasites lacking the endogenous FT2 gene. Together with an additional overexpression of tagged FT2 in wild type parasites, unaltered functionality of apicoplast-targeted FT2 is demonstrated and secondary effects can be excluded. Although fundamentally different localisation in the related Plasmodium species cannot formally be ruled out, this possibility is considered unlikely, not least because of the high sequence identity of the FT2 orthologs (86%). The re-evaluation of the physiological localisation in P. falciparum by episomal GFP fused FT2 expression was performed under the potentially overexpressing CRT promotor. Unsuccessful transfections, driven by secondary effects through overexpression in Pf3D7 cannot be excluded. On this account P. falciparum FT2 needs further localisation analysis to exclude a diverging localisation between the two Plasmodium species.
The apicoplast localisation of FT2 disqualifies this protein as a mediator of folate salvage. The other candidate for this molecular pathway, FT1, was shown to be dispensable for asexual blood stage growth in both rodent and human malaria models [87, 88]. We argue that Plasmodium folate salvage might not require a specialised uptake mechanism facilitated by plasma membrane transporters. Matz et al. have previously shown that the central folate metabolite salvaged by Plasmodium blood stages is the precursor pABA [74]. Owing to its aromatic benzene ring, pABA can diffuse across lipid membranes and thus access the cell passively [89]. The assumption of a plasma membrane folate transporter essential for Plasmodium blood stage development might thus be unwarranted. In transposon mutagenesis screens FT2 has been annotated with an intermediate phenotype in the asexual stages [87]. Therefore, we assess the unsuccessful FT2 deletion approaches in P. falciparum to be of technical reason instead of essentiality in blood stages. Together with previous biochemical observations [79], the distinct localisation of FT2 implies a role in internal trafficking like plastidial import or export of folates and/or related pteridine molecules.
67 3RD CHAPTER – FT2
Further biochemical assays, including metabolic profiling of purified apicoplasts [90] from WT and ft2– parasites are warranted to solve the molecular gatekeeper function of this Myzozoan-specific MFS transport protein.
Interestingly, FT2 does not contain a canonical bipartite leader sequence [91]. This motif, which consists of an ER signal peptide and an apicoplast transit peptide, is characteristic for proteins of the apicoplast stroma and of the innermost plastidial membrane [92-94]. By contrast, a transporter of the outermost apicoplast membrane, as well as three additional and yet uncharacterised apicoplast membrane transporters, lack this leader sequence [93, 95, 96]. The absence of a classical apicoplast targeting signal from FT2 might thus signify that FT2 is also inserted into one of the outer three membranes of the apicoplast. Additionally, FT2’s unexpected localisation illustrates - along with the previous examples - that screens for plastid targeting proteins may not be based on the respective targeting signals only.
Note that FT2 also localises to the apices of merozoites and sporozoites. Such a dual targeting pattern is not uncommon among Apicomplexans [86, 97]. Yet, the exact subcellular localisation of this fraction remains to be determined. However, it is interesting to speculate how this protein distribution might contribute to the defects in sporozoite formation and infectivity. We observed a striking correlation of the dual localisation during sporogony with the timing of life cycle arrest in FT2-deficient parasites. Whether the essential functions of FT2 in sporogony are promoted by the plastidial or extra-plastidial fraction, or both, remains unanswered at the moment. Absent effects in FT2-deficient schizonts imply that a potentially essential apical function of FT2 might at least be stage-dependent. However, the broad expression pattern of FT2 suggests that this transporter functions throughout the entire parasite life cycle, yet an essential role is only detected upon productive sporulation in the mosquito vector.
Counterintuitively to the broad expression pattern, a striking defect in FT2- deficient parasites was the vesiculation which is observed in oocysts only. How an imbalance in pteridine transport leads to this previously unrecognised phenotype remains hypothetical. Since this pattern is observed in a small proportion of late wild type oocysts as well, a metabolic imbalance leading to an increased growth-defect in oocyst is likely. The vital role of the apicoplast during blood infection is the biosynthesis of isoprenoid precursors as well as fatty acid synthesis and de novo heme biosynthesis
68 3RD CHAPTER – FT2
[6, 82, 98]. Recent work has established, that a loss of isoprenoid synthesis causes a halt in protein prenylation, which in turn results in defective vesicular trafficking [99]. In asexual blood stages, this causes fragmentation of the food vacuole and abnormal formation of the inner membrane complex (IMC). One could envisage insufficient production of isoprenoid precursors or other metabolites with vital roles in sporogony as a consequence of loss of FT2’s transport function, leading to membrane deformations, vesicles, swelling, and the general loss of infectivity in FT2-deficient sporozoites. However, a connection between FT2’s described cargo and isoprenoid precursor pathways is not obvious. Additionally, isoprenoid precursor associated defects would most likely be present in blood stages as well for which it was shown to be an essential metabolite [98, 100]. In contrast, type II fatty acid synthesis and de novo heme biosynthesis, two other important roles of the apicoplast, are not essential during the blood stage development in which the parasites draw on lipids and heme derived from the host [101-103]. Those factors change dramatically upon transmission to the insect host. However, in which way FT2’s described cargos could be involved in such pathways remains unclear.
One of the few evident folate-directed aspects in the apicoplast is the folate- based formylation of methionyl-tRNA by a methionyl-tRNA formyltransferase (FMT). FMT’s exclusive expression in liver stages, and no existent functional overlap with FT2 during life cycle progression suggests a plastidial role independent on an FT2 derived folate uptake and with no connection to the observed defect after the deletion of FT2. An additional described context of folate in the apicoplast is a potential impact on the apicoplast DNA replication. Purine synthesis is described as a critical folate-dependent pathway in rapidly growing cells [81]. Therefore, it is thinkable that during sporogony, which comes with an extreme multiplication of parasites and their apicoplasts, the development is impaired by an unfunctional or absent apicoplast in sporozoites. However, this latter aspect remains unaddressed in this study. As the functional study by Salcedo-Sora et al. focuses mainly on transport assays of folates, putative alternative cargos of FT2 have been neglected. For instance, the folate-biopterin family (FBT) is assessed as potential proton symporters [41]. Additionally, it is not guaranteed that FT2 transports specifically folates only. Promiscuity is a present phenomenon among MTPs [104]. Therefore, defects in FT2-deficient parasites can also be driven by co-transported molecules or unexpected cargos due to unpredictable promiscuity of the transporter. Such a scenario could explain, why a defect in the mosquito stages
69 3RD CHAPTER – FT2 is observed after FT2-deletion but not after interrupted de novo synthesis together with excluded salvage from the host [74].
Together, in the context of the current literature, the findings demonstrate the essential role of an apicoplast-resident folate transporter during Plasmodium sporogony and provide support for a functional link between FT2’s transport function and the production of the malaria parasite’s transmission stages.
70 4TH CHAPTER – PF3D7_1468600
4th Chapter: Role of a putative aminophospholipid ATPase (Pf3D7_1468600) with an ortholog loss in rodent malaria parasites 4.1 Introduction
In vitro culturing of P. falciparum and the rodent in vivo infection model in P. berghei are two well-studied malaria systems with differences in accessibility and relevance. Despite being two different organisms, the similarities between P. berghei and P. falciparum allow for bilateral studies that reveal insight about Plasmodium’s biology. Apparent differences between the two closely related species define the pathogen’s individuality and give insights about putative host- and therefore virulence- dependent features. Such potential virulence-associated characteristics are of particular interest for P. falciparum, the etiological agent of malaria tropica [7].
Remarkably, a range of genes can be found in Plasmodium organisms which are absent in rodent P. berghei parasites. The difference in genetic complexity is mainly discussed in the context of the host’s immune response or reproduction rate [105]. Beyond that, differences in MTP conservation between Plasmodium species seem reasonable in relation to the respective host-derived nutrients and their transport. In an amino-acid based reciprocal homology study, Weiner and Kooij identified MTP orthologues and their conservation among Plasmodium species and further organisms [31]. While most MTPs showed high conservation between Plasmodium species, a few P. falciparum MTPs were shown to have no orthologues in P. berghei. One example is a Pf3D7_1468600 encoded flippase which is a member of the P-type ATPases, a group of self-phosphorylating ion and lipid pumps. Based on similarities to conserved domains, the gene was assessed to encode for an aminophospholipid-transporting P- type ATPase [40]. Pf3D7_1468600 has been shown to be mainly expressed in ring stages and late gametocytes [146]. However, subcellular localisation and functional role have not yet been identified even though many P-type ATPases are associated with important functions in the parasite’s biology. This applies in particular to phospholipid flippases, of which the majority showed to be refractory to gene deletion in P. berghei [33]. Phospholipid flippases are restricted to eukaryotes and necessary to create phospholipid asymmetry, which is vital for several cellular processes [106]. Furthermore, the parasite consumes energy in the form of ATP, which suggests a crucial role of such ATPases for the parasite’s inner cost-benefit ratio. Which consequences the loss of this enigmatic flippase might have for P. falciparum was
71 4TH CHAPTER – PF3D7_1468600 investigated in this study by applying CRISPR/Cas9 reverse genetics on P. falciparum. Finally, the data allows to study a role of the P-type ATPase in the development of P. falciparum blood stage parasites.
4.2 Results 4.2.1 Phylogenetic analysis of Pf3D7_1468600
To confirm the loss of Pf3D7_1468600’s orthologs in P. berghei [31] and to extend this observation to other organisms with rodent hosts of the Vinckeia clade, an ortholog search was performed on PlasmoDB. Aligned with a phylogenetic assessment of the Plasmodium phylum [8], Pf3D7_1468600’s orthologs show to be present in the neighbouring and distant clades but not in the rodent parasitising Vinckeia clade (Fig. 14).
host organism gene (subgenus) Primates Plasmodium vivax PVX_117035 Plasmodium inui C922_00096 Plasmodium cynomolgi PCYB_124640 Plasmodium knowlesi PKNH_1212600 Plasmodium coatneyi PCOAH_00044220 Plasmodium fragile AK88_05106 Plasmodium ovale PocGH01_12046200 Plasmodium malariae PmUG01_12048300 Plasmodium berghei Rodents (Vinckeia) Plasmodium yoelii Plasmodium Plasmodium falciparum PF3D7_1468600 Primates (Laverania) Plasmodium reichenowi PRCDC_1467800 Plasmodium gaboni PGSY75_1468600 Plasmodium relictum PRELSG_1212200 Sauropsids Plasmodium gallinaceum PGAL8A_00228000
Fig. 14 Phylogenetic tree of Pf3D7_1468600. Indication of absence or presence of Pf3D7_1468600 and its orthologs in different Plasmodium organisms. Data shows loss of orthologs in Vinckeia clade (indicated by the red cross). Phylogenetic tree and branching of organisms based on [8].
4.2.2 CRISPR/Cas9-mediated disruption of Pf3D7_1468600
The generation of a Pf3D7_1468600 disrupted cell line was approached via a CRISPR/Cas9-mediated disruption design for Pf3D7_1468600 specific gRNAs expressed under a U6 promoter. Two constructs were designed with alternative gRNAs which induce a double-strand break at position 1640 and 1914 in the gene,
72 4TH CHAPTER – PF3D7_1468600 respectively. On- and off-target scores of the gRNAs are shown in Fig. 16 a. The respective gRNA targeting-sequences was annealed from ssDNA oligos with overhangs compatible to a digested BbsI restriction site and cloned downstream of a U6 promoter and upstream of a modified chimeric gRNA in the Cas9-U6 plasmid. For the donor template, homologous regions and a hDHFR drug cassette were assembled in a Gibson assembly to introduce all fragments in one step in the plasmid. The drug cassette served for positive selection after homology-directed repair of the nuclease induced double-strand break (Fig. 15 a and Fig. S 2). After transfection, cultivation under a constant WR99210 drug pressure led to positive transfectant populations after 33, 42 and 44 days for repeated transfection of the construct with a gRNA design for a double strand break at position 1914 (Fig. 16 a). While this gRNA resulted in three populations after three independent transfections, the alternative gRNA design (position 1640) was unsuccessful in six out of six transfections (Fig. 16 a). Transfected cultures were monitored for a period of at least 60 days. Transfectant populations were passed through two drug cycles (off-drug and on-drug period) until a stable population
(a) (b)
5‘WT 3‘WT
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Pf3D7_1468600 AmpR Cas9 gRNA disrupted - 1.0 kb
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Fig. 15 Experimental genetics approach for CRISPR/Cas9-mediated disruption of Pf3D7_1468600. (a) CRISPR/Cas9-mediated disruption strategy. Shown are the wild type (WT) locus (top), the transfection vector (middle) and the recombined locus (bottom). The targeting vector contains an Ampicillin resistance cassette for molecular cloning (AmpR, green), the Cas9 cassette (yellow) plus a guide-RNA (gRNA, red) with a gene-specific sequence. The donor template consists of a drug cassette for positive selection (hDHFR, blue) flanked by homologous regions (grey boxes) for a homology-directed repair serving for a partial deletion of the gene. Primer (P1-P6) annealing to wild type locus or inserted fragments allow diagnostic PCR to test for isogenic populations after drug- cycling. (b) Diagnostic PCR to test for wild type (WT) or integration specific (INT) locus indicated in
73 4TH CHAPTER – PF3D7_1468600
(a). Arrowheads show expected product size (details in Tab. S 2). Wild type samples (top) confirms correct amplification of wild type locus only. Pf3D7_1468600 disrupted cell line (bottom) indicates correct integration and absence of wild type locus.
(a) position 1640 position 1914
MTP MTP
off-target score1 49.9 45.7
on-target score2 70.3 66.8 1 2 3 transfected 6 transfections 42 33 44 population after unsuccessful days days days
(b) * *
1.0 W T t o grow th 0.5 ive Rela t
0.0 Pf3D7_1468600 WT disrupted
Fig. 16 (a) CRISPR/Cas9 transfections with two different gRNA positions on Pf3D7_1468600. Off-target and on-target calculation after 1: Hsu et al. [84] and 2: Doench et al. [85] gRNA at position 1640 showed no cell population after 60 days of culturing in six transfection repeats. gRNA at position 1914 showed cell populations between 33 and 44 days after transfection. (b) Relative growth of genetically disrupted Pf3D7_1468600 line compared to wild type growth measured by a SYBR-safe growth assay. SYBR-fluorescence output was adjusted to the background-noise and normalised to wild type signal. Shown is relative growth to wild type of measured cultures with mean and standard deviation. Mean relative growth rate is 0.927, while mean WT growth is adjusted to 1. Data are pooled from 4 replicates of 27 samples (total n=108 for Pf3D7_1468600 disrupted, total n=107 for Pf3D7 wild type). **, P<0,01; Student’s t-test on means of each replicate assuming Gaussian distribution.
74 4TH CHAPTER – PF3D7_1468600 was re-established. For the studied cultures, this happened within two asexual cycles. Genomic DNA of the transfectant lines was tested with specific primer combinations (P1-P6) indicated in Fig. 15 a, to test correct integration and absence of the wild type locus. Diagnostic PCR was performed on cell populations from two independent transfections. Recombination specific PCR bands (5’INT and 3’INT) and the absence of wild type parasites can be identified in the disrupted cell line, while wild type DNA confirms 5’ and 3’WT products and no product for integration specific primer combinations (Fig. 15 b). One transfectant strain displays an unexpected band in the 5’ integration product (5’INT) and was therefore excluded. Further analysis was performed on the cell line which was positive for diagnostic PCR only (Fig. S 1).
4.2.3 SYBR-safe growth assay indicates that Pf3D7_1468600 is involved in growth-efficiency in the asexual blood stages
To test the growth rate of parasites with a genetic disruption of Pf3D7_1468600, a growth assay was performed in a comparable manner to SYBR-safe drug assays (see 2.2.7 in methods). For the Pf3D7_1468600 disrupted cell line, four replicates of 27 wells of synchronised parasites were incubated for 72 hours in 96-well plates in parallel with wild type parasites and uninfected RBC background controls. SYBR-safe exposure and plate reader measurements at 490nm excitation/520nm emission settings provide a fluorescence intensity which serves as measure for DNA quantities after offsetting background signal from the uninfected erythrocyte controls. The fluorescence output of each well of the Pf3D7_1468600 disrupted line was divided by the mean wild type output of the same 96-well plate. The quotient reflects the relative growth of the disrupted cell line compared to wild type parasites. The ratio of 0.927 is equivalent to a 92.7% growth rate compared to wild type development in the asexual stages for Pf3D7_1468600 disrupted parasites (Fig. 16 b).
75 4TH CHAPTER – PF3D7_1468600
4.3 Discussion
By piggyBac transposon mutagenesis Pf3D7_1468600 was preassessed as not essential but with effects on the fitness of the mutant [87]. The data of this study support this evaluation by successful interruption of Pf3D7_1468600 combined with a growth defect of the interrupted cell line. The defect, with ~7.3% less growth compared to equally synchronized WT parasites identified by SYBR-safe measurements, is based on a non-continuous culture with a 72-hour incubation. Long-term growth defects could potentially be identified by competition assays between disrupted cell lines and WT parasites in pooled cultures. The assessment of outcompeting wild type growth could be analysed by Pf3D7_1468600-directed qPCR. 7.3% less parasites after 72 hours indicates that wild type parasites might outcompete Pf3D7_1468600- disruted cell lines after several weeks of competitive co-culturing, when compensatory growth effects are left out of consideration. Therefore, the described short-term defect already suggests a growth-related function in asexual stages, hypothetically based on aminophospholipid-imbalance. In addition, previous transcriptome studies on flow cytometry sorted gametocytes show an upregulation of Pf3D7_1468600 in male gametocytes [107]. Therefore, a role during gametocytogenesis of male gametocytes would be reasonable. Further assessment in gametocyte culture could include male:female ratios or analysis of different lipid-class patterns like in [108]. Exflagellation assays, like in [109], would indicate if the transporter has a role during the drastic membranous reorganisation of male gametocytes. Beyond that, infection experiments of the mosquito vector would allow to asses defects during host- transmission. As the experiments were performed in vitro only, it is not clear to which extend the adjusted culturing conditions mask a role of Pf3D7_1468600. It would be worth it to investigate, if the observed effects enhance through a reduction of potential substrates.
In this study, no functional characterisation was performed except for the growth assay. However, the measured defect shows that even though the MTP is not essential, it is linked to P. falciparum’s development in asexual stages. An absence of this function in P. berghei might be therefore related to (i) differences in the cell biology of the asexual stages which make the function of Pf3D7_1468600 not affordable for the cost of ATP in P. berghei, (ii) different transport-requirements due to differences in the composition of host-derived substrates, or (iii) a functional impact of Pf3D7_1468600 related to the transmission from vertebrate to mosquito which is not
76 4TH CHAPTER – PF3D7_1468600 required in rodent malaria. Exemplified for the latter aspect, for some genes which are absent in rodent parasites, a role in the gametocytogenesis of P. falciparum was described [105]. This could be in particular comprehensible since parameters which influence a successful transmission from host to mosquito might vary between the respective host species. But such conclusions remain speculative and are not supported by the shown data.
Transfections of P. falciparum have been done for many decades [43] and were revolutionised by the possible application of CRISPR/Cas9 genome editing [46, 47]. Through the active induction of a double-strand break, genome editing can exploit the cellular repair machinery with higher efficiencies compared to conventional double homologous recombination [110, 111]. However, P. falciparum has an extremely (A + T)-rich genomic sequence with particular low GC-content in intergenic regions [57]. The (A + T)-rich genome limits, first of all, flexibility in the design of -NGG-nucleotide protospacer adjacent motifs (PAM), and secondly, finding appropriate annealing regions for the primer design for homologous regions outside of the ORF. Furthermore, it is recommended to keep the distance of homologous regions from the double-strand break rather short, which limits the size of deleted fragments [112]. To address this, two distant gRNAs can be co-transfected simultaneously, whose distant annealing sites define the size of the deleted fragment. Nevertheless, big differences in gRNA- efficiencies are reflected in the results of this study, showing either 100% or 0% successful recombination of the respective gRNA design (Fig. 16 a). Therefore, a design with two co-transfected gRNAs requires a previous experimental check on the gRNA efficiency. A correlation between gRNA efficiency and designed on-target scores is not apparent in the performed experiments.
With the described construct design, only genetic disruption with a partial deletion of the gene can be achieved. In the ~6.2 kb gene a ~700 bp fragment was replaced with the ~1.8 kb drug cassette. Diagnostic PCR proves the replacement of the gene fragment with the drug cassette but does not indicate if the deletion leads to a loss of Pf3D7_1468600’s function. Analysis on transcript and protein level could indicate such downstream effects, which was not performed in this study. Additionally, off-target efficiencies were optimised during the design, but a complete exclusion of off-targets is not guaranteed. But so far, off-target lesions have neither been reported nor systematically examined after CRISPR editing in Plasmodium [110].
77 4TH CHAPTER – PF3D7_1468600
Constant drug pressure after transfection led to rather late-appearing transgenic populations compared to other studies [47], in which drug pressure is often removed a few days after transfection. In this set-up CRISPR/Cas9 transfected populations do not develop faster (33 – 42 days) than conventional knock out strategies which rely on rare homology-driven integration events. It may be speculated whether the slow growth is connected to the growth phenotype of gene disruption. Diagnostic PCR proved an absence of WT parasites immediately after drug cycling. Therefore, it can be assumed, that constant drug pressure after transfection circumvents time-consuming clonal isolation since populations carrying the aimed modification are already isogenic. Still, drug-cycling is necessary to exclude parasites carrying an episomal plasmid. Since the populations recovered almost immediately after drug cycling, recombination via double homologous recombination from the donor region of the CRISPR/Cas9 plasmid seems unlikely. Additionally, such recombination would have occurred in plasmids carrying the 1640 gRNA site as well. Therefore, successful genetic disruption of Pf3D7_1468600 via CRISPR/Cas9-mediated recombination can be assumed.
In summary, the successful CRISPR/Cas9-mediated genetic disruption of Pf3D7_1468600 suggests a minor growth defect in asexual stages. This putative aminophospholipid transport is not essential in P. falciparum and got lost during phylogenesis in rodent malaria parasites. To understand a functional context of the role in P. falciparum, a functional characterisation, further investigations on gametocytogenesis and an identification of Pf3D7_1468600‘s localisation are necessary. A complete deletion of Pf3D7_1468600 by a co-transfection of two distant gRNAs would be necessary to confirm the results.
78 5TH CHAPTER – CRT AND ATP4
5th Chapter: Expression profiling of two vital P. berghei membrane transport proteins, CRT and ATP4
Among many undescribed MTPs, the literature pays particular attention to two candidates with gaining prominence, due to their associations with drug resistance and drug susceptibility: The Chloroquine Resistance Transporter (CRT, Pf3D7_0709000 and PBANKA_1219500), in which mutations in the molecule can confer chloroquine- resistance to the parasite, and the cation-ATPase ATP4 (Pf3D7_1211900 and PBANKA_0610400), which is a promising drug target inhibited by spiroindolones. For both transporters, substrates are extensively discussed. However, functional assignments of MTPs in P. falciparum in vitro studies are often limited to the experimental examination of blood stages. Nevertheless, an important step towards a functional classification is the determination of the expression during life cycle progression and their intra-cellular localisation.
In the P. berghei infection model, the entire Plasmodium life cycle can be accessed both in vivo and in vitro. Combined with the application of experimental genetics, this chapter addresses the determination of the spatio-temporal expression pattern of the two essential MTPs during the entire life cycle progression in P. berghei. Live fluorescence microscopy of transgenic P. berghei parasite lines that harbour endogenously tagged MTP::mCherry permits a systematic analysis in comparison to normal wild type parasites via digital image processing with ImageJ. Signal quantification is used to identify down- and up-regulations during the parasite’s in vivo development as a step towards further functional classification of CRT (in section 5.1) and ATP4 (in section 5.2).
5.1 The Chloroquine Resistance Transporter 5.1.1 Introduction
Chloroquine (CQ) represents one of the most successful antimalarial drugs in the last 90 years of antimalarial research. It is active against all Plasmodium species and has low side effects, high stability, is simple to apply and cheap in the production [113]. Therefore, CQ became indispensable for malaria control [114]. However, resistances emerged from several origins and spread throughout malaria-endemic regions, leading to severe increases in the mortality of the disease [115-119]. As a potential mechanism
79 5TH CHAPTER – CRT AND ATP4 of resistance, the extrusion of CQ which leads to less CQ accumulating in the parasite, was observed [120, 121]. This process is governed by a ~330 kDa protein with polymorphisms linked to the resistance [37, 122, 123]. In the resistant protein, lysine 76 (K76) appears to be replaced by the uncharged amino acid threonine (K76T) [124, 125]. The protein was identified to be a 10 trans-membrane domain MTP of the digestive vacuole (DV) and carries the name Chloroquine Resistance Transporter (CRT), independent of harbouring the resistance transmitting variation or not [37]. Later it was shown that mutant CRT expressed in Xenopus laevis oocytes transports CQ, while the WT protein does not [126]. One hypothesis behind this resistance- conferring modification in CRT is as follows: Neutral CQ gets di-protonated and accumulates in the acidified DV. In resistant parasites, the loss of a positive charge (K76T) initiates drastic changes in the electrostatic potential of CRT, and di-protonated CQ can bind to a predicted binding site and be exported from the DV [127, 128]. However, K76T is assumed to be critical for drug resistance, but additional point mutations are relevant for CQ export efficiency [126, 129]. Similar to CQ resistances, reduced drug effectivity, induced by a distinct set of point mutations in CRT, has recently been associated with piperaquine (PPQ) resistances [130].
CQ’s generally acknowledged mechanism of action is described as follows: Inside of the red blood cell, the parasite ingests haemoglobin, which is delivered to the acidic DV and proteolyzed. Toxic heme results as a by-product, which is detoxified through the polymerisation to crystalline b-hematin, known as hemozoin [28]. CQ can diffuse across membranes and accumulates in the digestive vacuole [131], where it potentially inhibits the detoxification of heme [132]. The mechanism of the inhibition is hypothesised to be due to the formation of CQ-hemozoin complexes [133-135]. In recent years, a potential interaction of CQ with CRT and a potential inhibition of its native function became an alternative approach to interpret the activity [132].
Even though decades of research have dissected the background on CQ resistances and CRT, the native function is so far not yet entirely revealed. Disruption of the gene was shown to be deleterious for parasite viability, and P. falciparum CRTs function is likely to be essential [136]. Remarkably, after discontinuation of CQ use, a long-term decline in the frequency of parasites with mutant CRT was shown in field studies [137]. Additionally, lower parasite densities and higher gametocyte conversion rates are observed for the mutant alleles [138]. Such results describe the fitness cost of mutations in CRT and underline the importance of its native function. The
80 5TH CHAPTER – CRT AND ATP4 identification of the physiological substrate is still pending, and is so far predicted to be haemoglobin-derived peptides or amino acids. Such predictions are mainly based on CRT’s localisation, its classification as a drug/metabolite transporter superfamily (DMT) member, as well as metabolite studies [126, 139-141]. Furthermore, it was described to be a proton-coupled transporter recognising cationic substrates [142] and showing electrogenic iron transport when expressed in Xenopus oocytes [143]. However, the physiological role of the transport, the transport mechanism and the functional regions of CRT are not fully characterised. Based on DMT templates with shared structural features, CRT’s tertiary structure, its functional domains, and exporter-properties can be predicted [128, 144]. Structural modelling allows the identification of putative binding pockets which cut down the list of potential substrates [128].
Even though CRT has been the focus of different scientific disciplines for many decades, the majority of the studies is dedicated to either resistance-linked properties or substrate identification. In this context, experimental genetics has mainly been used on CRT as an instrument for functional assessments to confirm a relation between point mutations and CQ resistance. Beyond that, experimental genetics has so far been a powerful tool which revealed insight about CRTs essentiality [136] and the mutant’s selective advantage in transmission to the mosquito [145]. Even though CRT’s functional role is understood to be mainly condensed to trophic blood stages, an extended expression along life cycle progression including mosquito stages [146] and early ring stages [147] indicates a role beyond the trophic phase of the parasite’s development. In this study, transgenic parasites with an endogenously tagged CRT::mCherry are used to determine a spatio-temporal expression pattern of CRT during the entire P. berghei life cycle progression. An expression beyond the blood stage parasites, could suggest additional functions of CRT in the development of Plasmodium parasites.
5.1.2 Results CRT 5.1.2.1 An isogenic CRT::mCherry cell line was used for spatio-temporal analysis
S. Kenthirapalan has previously generated the CRT::mCherry cell line. The carboxy-terminus was cloned in frame with an mCherry-3xMyc tag and a 3’ flanking region distal of the drug cassette (Fig. 17 a). The C-terminus and the 3’ flanking region allowed the integration of a fluorescent tag, GFP- and drug-cassette via double
81 5TH CHAPTER – CRT AND ATP4 homologous recombination. Flow cytometric isolation of transfected parasites enables the generation of an isogenic population. Diagnostic PCR with specific primers producing a PCR product for correct assemblies confirms the endogenous integration (5’INT and 3’INT) and proves the absence of WT parasites (3’WT, Fig. 17 b).
(a) 5‘WT 3‘WT (b) WT INT P1 P2 P3 P4 CRT 5‘ 3‘ 5‘ 3‘ - 5.0 kb
ANKA - 1.5 kb P5 P6 tag GFP cassette drug cassette - 0.5 kb - 5.0 kb
CRT::mCherry - 1.5 kb 5‘INT 3‘INT
P1 P5 P6 P4 - 0.5 kb CRT tag GFP cassette drug cassette
Fig. 17 Experimental genetics approach for endogenous tagging of CRT. (a) The targeting plasmid contains an mCherry-3xMyc tag (red), the CRT carboxy-terminus and the 3’ flanking regions (grey boxes) to generate an in-frame carboxy-terminal mCherry-3xMyc (tag, red) fusion protein. Shown are the wild type (WT) locus (top), the transfection vector (middle) and the recombined locus (bottom). The plasmid also contains a GFP- and a drug cassette (green and blue). Construct design, cloning and generation of isogenic CRT::tag cell line was done by S. Kenthirapalan. P1-P6, primer annealing sites used for diagnostic PCR to prove absence of wild type (WT) and presence of integration (INT)-specific templates. (b) diagnostic PCR with primer combinations from (a) confirm absence of WT parasites in the recombinant CRT::mCherry parasite line. Arrowheads show expected product size (details in Tab. S 1).
5.1.2.2 Food vacuole resident CRT::mCherry co-localises with hemozoin crystals.
In differential interference contrast microscopy (DIC), the hemozoin crystals appear in high-contrast reflections. A merge of DIC and mCherry signal shows a noticeable correlation of hemozoin structures with the food vacuole localising CRT::mCherry signal in different depths of fixed gametocytes (Fig. 18). A comparable distribution of CRT::mCherry can be observed for other blood stages, like trophozoites, schizonts and ookinetes (Fig. 19 a and b). Note that in particular in trophozoites, an accumulation of CRT::mCherry at the cell periphery is observable (Fig. 19 a, white arrowhead). In schizonts, the CRT carrying food vacuole appears in one centralised compartment, the so-called residual body, surrounded by newly formed merozoites. In gametocytes and ookinetes, the signal scatters in the cell lumen in compartments of differed sizes.
82 5TH CHAPTER – CRT AND ATP4
The post erythrocytic stages, which harbour no crystalloid of hemozoin, express CRT::mCherry as well. However, a reliable identification of the localisation is limited by fluorescence microscopy. Oocysts show a rather low-level expression, which remains unchanged during internal sporoblast formation and sporozoite development (Fig. 19 b). In both, intra-oocystic sporozoites as well as sporozoites extracted from the mosquito’s salivary glands, CRT::mCherry shows no distinct expression (Fig. 19 c). 48h after Huh7 infection, a potentially new localisation associated with cytosol, the PPM and/or PVM can be postulated in developing EEFs. For later EEF stages CRT::mCherry’s signal becomes less distinct.
Overall, CRT localises in the digestive vacuole in trophic stages. Later stages show a carry-over of the lysosomal compartment up to ookinetes. Furthermore, EEFs show an expression, but with indistinct localisation. The signal in oocysts is not distinguishable from background fluorescence (compare Fig. S 4).
Fig. 18 CRT::mCherry fluorescence signal co-localises with hemozoin crystals in gametocytes. Shown are two DIC and mCherry-fluorescence images of the same PFA-fixed gametocyte at different focal planes. The red mCherry signal co-localises with high-contrast hemozoin crystals. Bar: 10 µm.
83 5TH CHAPTER – CRT AND ATP4
(a) DIC CRT::tag (b) DIC CRT::tag CRT::tag CRT::tag cytoplasm DAPI cytoplasm DAPI
ookinete
ring stage
oocyst d10
trophozoite blood stages
schizont
oocyst d14 ookinetes and mosquito stages
oocyst d17
gametocyte
(c) DIC CRT::tag CRT::tag cytoplasm DAPI
sporozoite
EEF 24h sporozoites and EEFs
EEF 48h
EEF 69h
Fig. 19 Spatio-temporal expression of CRT::mCherry. Fluorescence signal of CRT::mCherry expressing parasites imaged throughout all stages of the P. berghei lifecycle (a-c). Shown are representative fluorescent and light microscopy images of the respective stages: (a) sexual and asexual blood stages. White arrowhead indicates CRT coming from the parasites periphery; (b) ookinetes and mosquito midgut oocysts; (c) sporozoites and 24h, 48h and 69h EEFs after Huh7 infection with sporozoites. Merge of differential interference contrast images (DIC) and cytosolic expressed GFP (green) for parasite identification (1st column); merge
84 5TH CHAPTER – CRT AND ATP4 of CRT::mCherry signal (red) with Hoechst 33342 nuclear stain (DAPI, blue, 2nd column); CRT::mCherry signal in greyscale (3rd column). Bar 10 µm.
5.1.2.3 CRT::mCherry shows an expression pattern beyond blood stages.
To assess an expression quantification for CRT during life cycle progression, live cell fluorescence microscopy was performed on all life stages under equal exposure times. Measurement of mCherry-fluorescent intensity output of a whole parasite was adjusted to background noise and WT autofluorescence of the respective parasite stage and used to calculate signal density/µm2 (described in methods 2.2.2). The quantified output allows the assessment of a stage-specific CRT::mCherry expression and the recognition of a life cycle progression dependent expression pattern of the protein.
Blood stages show different expression levels of CRT::mCherry for the asexual cycle and the sexual stages (Fig. 20 a). In the asexual cycle, CRT expression is low in ring stages and reaches its expression peak in trophozoites. Schizonts from overnight cultures display a lower expression compared to trophozoites. For the parasites entering sexual differentiation, the increase from ring stages to trophozoites continues for gametocytes, which represent the stage with the highest CRT expression within the entire life cycle progression. In ookinete cultures, fertilised female gametocytes differentiate to motile ookinetes, which still show a substantial amount of CRT::mCherry fluorescence intensity, comparable to quantities expressed in trophozoites. After the transmission of the parasites from mouse to mosquito, developing oocysts can be observed in the basal lamina after mosquito midgut preparation (Fig. 19 b). Autofluorescence for midgut stages was remarkably higher, which leads to an increase in variance of measurements taken for each developmental stage of the oocysts. Due to the autofluorescence adjustments with WT oocysts, data points are levelled and can reach negative values (compare autofluorescence of wild type parasites in Fig. S 4). The mean values of the different oocyst’s stages fluctuate around zero. Intra-oocystic sporozoites had no impact on the measurements of late oocysts. The absence of a fluorescence signal in salivary gland extracted sporozoites confirms this. Parasite infecting liver cells show increasing expression of CRT::mCherry, starting with a low signal in 24h p. i., which increases at 48h and slightly decreases at 69h before the merosomes emerge from the infected liver cells. In general, late EEFs show a data variation comparable to blood stages.
85 5TH CHAPTER – CRT AND ATP4
Overall, CRT::mCherry fluorescence intensity is strongest in blood stages, with peak intensity in gametocytes. Beyond that, the measurements reveal an additional CRT::mCherry expression in late liver stages.
(a) (b)
CRT-mCherry CRT qPCR
1000 1 ± SD 2 500 SP 70 /1) 0.1 H m expression
0 0.01 Int. Density / (normalised to relative mRN A -500 0.001
oocysts ringstages schizonts ookinetes EEFsEEFs 24h EEFs48h 69h schizonts ookinetes EEFsEEFs 24h 48h sporozoites sporozoites EEFs 72h trophozoitesgametocytesoocystsoocysts d10oocysts d14 d17 blood stagesgametocytes
Fig. 20 Stage specific expression profiling of CRT during life cycle progression. (a) integrated CRT::mCherry live fluorescence density/µm2 analysed for parasites with mean (line) and standard deviation (error bars) of respective life stage. (b) relative CRT mRNA expression measured by qPCR on cDNA converted from RNA of respective life stage (provided by K. Müller). Bars show mean and standard deviation. qPCR was performed in triplets except for sporozoites (one replicate) and EEF 24h (two replicates). qPCR has been normalised to the housekeeping gene HSP70/1.
86 5TH CHAPTER – CRT AND ATP4
5.1.2.4 Validation of CRT::mCherry signal quantification by stage-specific mRNA expression profiling
In this approach, we want to align the results of the CRT::mCherry expression fluorescence measurements with a standard technique for expression analysis. The wild type cDNA was provided by K. Müller. The relative CRT mRNA steady-state levels are normalised to the transcript levels of the housekeeping gene HSP70/1 on wild type parasites. The overall CRT transcription is much lower compared to HSP70/1, which is reflected in quite low relative mRNA expression values (Fig. 20 b). Since mRNA extraction cannot be done separately on ring stages and trophozoites, expression from mixed blood stages represents an average expression of ring stages, trophozoites, and gametocytes. Therefore, CRT mRNA expression is potentially increasing from ring stages and trophozoite to gametocytes, since mRNA from gametocyte cultures shows an even higher expression than mixed stages. A more substantial increase can be observed from mixed stages to schizonts, which represent the stage with the highest expression among all life stages. In ookinetes, the expression drops, and becomes negligible in oocysts and sporozoites. Liver stages show a comparable expression to blood stages with an expression peak at 48h.
5.1.3 Discussion CRT
This is, to our knowledge, the first full life cycle P. berghei CRT-expression analysis with simultaneous confirmation of the localisation. The data from both qPCR and CRT::mCherry intensity measurements reveal an expression of CRT beyond blood stages, suggesting additional functions in other life stages. Previous transcriptomic studies showed that CRT expression is not exclusively tied to the trophic stages. Illumina-based RNA sequencing showed an increasing CRT expression from ring stages to trophozoites and a lower expression in schizonts and gametocytes [148]. RNA sequencing of sorted gametocytes confirmed an expression decrease from erythrocytic stages to gametocytes, in which the expression is predominantly driven by the male gametocytes [149]. Unpublished transcriptional data by Hoejmakers (accessible on PlasmoDB) suggest a continuous decline of CRT expression from rings to schizonts in Pf3D7. It is not possible to distinguish between rings stages, trophozoites and gametocytes in the obtained blood stage sample in our study. Therefore the average blood stage expression is inconsistent with the previously
87 5TH CHAPTER – CRT AND ATP4 described transcription data, by showing slightly lower qPCR-expression compared to schizont and gametocyte cultures. Variation between the set-up of the biological samples (e.g. in vivo blood stages vs. in vitro schizont and ookinete stages), between different studies as well as different transcriptomic techniques might have an impact on the comparability of such results.
On the protein level, the CRT::mCherry expression peak in trophozoites links a functional association of CRT to the trophic phase in asexual parasites, which is consistent with previous data shown for the P. falciparum CRT [150]. Having said this, the data suggests the strongest P. berghei CRT::mCherry expression in gametocytes. CQ kills early gametocytes but has no effect on later gametocyte stages, which suggests haemoglobin metabolization to be essential up to early gametocytes [151]. It is described that the parasite takes up far more haemoglobin than it needs for its own metabolism [152]. Therefore, the DV and its content could be carried over in late sexual stages. Late gametocytes could potentially draw on haemoglobin-derived metabolites from previous metabolization which would potentially maintain CRTs putative export- function. Another explanation for higher CRT::mCherry expression in gametocytes than in trophozoites, could be a higher likelihood of trophozoites with a high CRT abundance to enter gametocytogenesis. However, most of the discussed triggers for gametocytogenesis are understood to favour an escape from an unfavourable environment instead of being ‘well fed’ [19]. By the conversion to zygotes and ookinetes, an uptake of erythrocytic haemoglobin can be ultimately excluded. Thus, it appears that, CRT’s export function is either obsolete or based on stored metabolites in the DV. However, since the parasites remodelling during ookinete conversion is not entirely understood [153], carryover of the DV cannot be distinguished from a functional need of the DV and CRT. The qPCR shows diminished but not absent CRT mRNA expression which supports the latter suggestion. An interpretation of the CRT::mCherry expression data in the mosquito stages is limited due to high variation and strong autofluorescence of oocysts imaged on midguts. Nevertheless, qPCR data indicates that CRT is poorly expressed in oocysts. Consistent with that, sporozoites show neither CRT::mCherry nor strong qPCR expression. Strikingly, the additional expression in EEFs could give new hints about its native function. Precisely because no haemoglobin uptake occurs in EEFs, the observed expression reveals either a potential new function in hepatic stages, or an early expression during the formation of hepatic merozoites. Since a clear localisation cannot be identified, functional
88 5TH CHAPTER – CRT AND ATP4 implications remain speculative. However, if CRT has a functional role in EEFs it could reject a hypothetical inhibition of CRTs native function by CQ [132], since CQ shows no, or only marginal effects on hepatic stages [154, 155]. But without any further evidence about CRTs function and localisation in EEFs, a membranous or transient expression in the cytosol for emerging merozoites seems most likely, and is consistent with PPM association in later ring stages.
The association of CRT and the DV in the blood stages is sufficiently understood and CRT::mCherry can serve as a DV marker [127] to monitor DVs morphological changes. The digestive vacuole gets formed through micropinocytosis to take up haemoglobin from the host cell starting in early ring stages until gametocytes [156, 157]. The pinocytosing vesicles are coming from the parasite’s periphery. Interestingly, the data show, that CRT::mCherry is already partly associated with the parasite’s membranes and/or pinocytosis events in trophozoite and potentially ring stages (Fig. 19 a, white arrowhead). Therefore, it is likely, that CRT::mCherry is already localised in the parasite’s plasma membrane. An impact of CRT on pinocytosis regulation cannot be excluded. In the pinocytosed vesicles, the now inner PVM gets degraded, which implies CRT’s potential expression in the PPM [158]. Smaller vesicles fuse together and form a bigger digestive vacuole in trophozoites and schizonts before they scatter up in multiple smaller DVs in gametocytes and ookinetes. In liver stages, a DV does not exist and a signal localisation cannot be distinguished accurately. It is likely, that the signal is either cytosolic or membrane-associated for a later localisation in merozoite-membranes, similar to the observed localisation in early blood stages as described above.
In total, the CRT::mCherry analysis provides insights into CRT’s expression pattern and therefore it’s potential functional relevance. Besides the well-known expression in blood stages, strong expression in gametocytes and ookinetes have been identified. The expression in liver stages gives hints about either a potential new function or an early expression in liver merozoites in preparation for the intra- erythrocytic trophic phase. At the end, even though CRT has been studied for many decades, fundamental details of CRT’s characteristics remain enigmatic.
89 5TH CHAPTER – CRT AND ATP4
5.2 ATP4 5.2.1 Introduction
ATP4 is a P-type ATPase, with a broad range of potential inhibitors [159, 160]. It has previously been described to be a Ca2+-ATPase of apicomplexan organisms [161]. Later, numerous studies provided adequate evidence that ATP4 is a Na+-efflux ATPase which exports Na+ against the gradient. This was in particular shown through the use of ATP4-targeting antimalarials and their impact on the parasites cell physiology.
ATP4 was suggested as a target for spiroindolones, a safe and potent group of new antimalarials that kill blood stage parasites in nanomolar concentrations [162]. Such spiroindolones - and a big range of other ATP4 directed drugs - interfere with the parasite’s Na+ homeostasis [42, 160] and the Na+-dependent ATPase activity in the parasite [36, 163]. After concentration-dependent spiroindolone application, Na+- content increases proportionally [164, 165]. The drug-driven Na+-accumulation in the parasite results in a Na+-dependent swelling, contributing to the inhibition of parasite growth [166]. In agreement with that, the compound loses its efficacy when the extracellular Na+ concentration is reduced [167]. Mutations in ATP4, as they have been profiled by Rottmann et al. [162], can confer resistances against compounds like spiroindolones and other Na+ homeostasis interfering drugs. In return, transgenic parasites with an introduction of the resistance-associated mutations display reduced sensitivity to growth inhibition by the compound [162, 164, 167]. Interestingly, mutant
+ + ATP4 showed increased resting of [Na ]i, decreased Na efflux, and a higher sensitivity to growth-inhibitory effects of excessive extracellular Na+ exposure [42]. Therefore, the fitness cost of resistant parasites could suppress a selection of resistant strains [35]. Together with its fast-acting antimalarial activity, drugs active against ATP4 are a promising new drug-group with a few candidates in clinical trials (KAE609 and (+)- SJ733). Additionally, the diverse range of drugs active against ATP4 indicates that the Na+ pump is a potential ‘Achilles heel’ of the parasite. However, the actual mechanism by which such drugs inhibit ATP4’s native function is not clear yet.
It was postulated that ATP4 extrudes Na+ in exchange for H+, which imposes an acid load on the parasite. Such acidification was observed to be dependent on extracellular Na+ [168]. Consistent with this, inhibition of ATP4 resulted in a cytosolic alkalisation [42]. Consequently, a pH dependent regulation is discussed [163].
90 5TH CHAPTER – CRT AND ATP4
Like in all other cells, the parasite maintains a low Na+ concentration in the cytosol. Uninfected RBCs counter act the electrochemically driven influx of Na+ by utilising Na+, K+ ATPases to maintain a low internal Na+-concentration. After an invasion of the parasite, the red blood cell undergoes drastic changes in its ionic composition [169]. Such changes are driven by the induction of new permeability pathways (NPP) in the RBCs which alter the permeability of the RBC’s membrane. Through channels being located in the RBCs plasma membrane, the increased permeability enables the exchange of nutrients and “waste” products as well as low molecular weight solutes including ions [170]. The induction takes place first at 18 hours after infection, and Na+ influx and K + efflux occur [171, 172]. The activity of RBCs Na+, K+ ATPases increases temporarily to counteract the uncontrolled ion changes, but it is not efficient enough to successfully maintain a required equilibrium [171]. Interestingly, the induced Na+ increase in the erythrocyte’s cytosol is described to be not essential for the parasite’s development [173]. Following the same pressures as the RBCs, intraerythrocytic parasites are suggested to maintain a low cytosolic Na+ concentration by extruding Na+ with ATP4 [42, 174]. Membrane ATPase assays suggest, that the majority of the parasite’s total Na+ dependent ATPase activity is contributed by ATP4 [163]. This ATP4-associated ATPase activity increases with increasing Na+ concentrations [42, 165], and can be upregulated up to 3.2-fold [163]. Consistent with the assigned function, it was shown that ATP4 localises in the PPM by the generation of a P. falciparum ATP4-GFP fusion protein [162]. How, and in which extend the inward Na+ flux occurs via the PPM, is not entirely clear [172]. At the same time, it is unclear why low cytosolic Na+ concentrations are of cell-physiological importance for the parasite’s development [175].
Due to the fast clearance of parasites after drug application, ATP4’s function can be assumed to be highly essential. Loss of P. falciparum ATP4 has shown to be deleterious to blood stage parasite growth [176]. Nevertheless, it is not known if the drastic environmental changes that occur throughout the rest of the parasite life cycle requires functional ATP4 due to the potentially different ion concentration in mosquito and liver cells. Furthermore, a profiling of ion-compositions in mosquito midgut, basal lamina, mosquito’s haemocoel, saliva and finally liver cells requires further studies.
In a similar methodological approach to CRT, we apply experimental genetics to determine the expression intensities of ATP4 and its intracellular localisation during the entire P. berghei life cycle progression. Expression intensities are digitally
91 5TH CHAPTER – CRT AND ATP4 processed based on live fluorescence microscopy of transgenic P. berghei-parasite lines that harbour endogenously tagged ATP4::mCherry. Together with qPCR transcription analysis, potential functions of ATP4 will be assessed based on the expression beyond blood stage parasites, always in strict coherence with the respective localisation.
5.2.2 Results ATP4 5.2.2.1 An isogenic ATP4::mCherry cell line was used for spatio-temporal analysis
Design and generation of parasites are similar to the description for CRT (Fig. 21 a) and were previously generated by S. Kenthirapalan. To check for correct integration via double homologous recombination and the absence of WT parasites in the isogenic cell-line, diagnostic PCR was performed on genomic DNA. Primers producing a PCR product specific for correct assemblies, confirm the endogenous integration (5’INT and 3’INT). Primer combinations suitable for the genomic wild type locus proves the absence of wild type parasites in the isogenic cell line (3’WT, Fig. 21 b).
(a) 5‘WT 3‘WT (b) WT INT P1 P2 P3 P4 ATP4 5‘ 3‘ 5‘ 3‘ - 5.0 kb
ANKA - 1.5 kb P5 P6 tag GFP cassette drug cassette - 0.5 kb - 5.0 kb
- 1.5 kb 5‘INT 3‘INT ATP4::mCherry
P1 P5 P6 P4 - 0.5 kb ATP4 tag GFP cassette drug cassette
Fig. 21 Experimental genetics approach for endogenous tagging of ATP4. (a) The targeting plasmid contains an mCherry-3xMyc tag (red), the ATP4 carboxy-terminus and the 3’ flanking regions (grey boxes) to generate an in-frame carboxy-terminal mCherry-3xMyc (tag, red) fusion protein. The plasmid also contains a GFP- and a drug cassette (green and blue). Construct design, cloning and generation of isogenic ATP4::tag cell line was done by S. Kenthirapalan. P1-P6, primer annealing sites used for diagnostic PCR to prove absence of wild type (WT) and integration (INT)-specific templates. (b) diagnostic PCR with primer combinations from (a) confirm absence of WT parasites in the recombinant ATP4::mCherry parasite line. Arrowheads show expected product size (details in Tab. S 1).
92 5TH CHAPTER – CRT AND ATP4
(a) DIC ATP4::tag (b) DIC ATP4::tag ATP4::tag ATP4::tag cytoplasm DAPI cytoplasm DAPI
ookinete
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(c) DIC ATP4::tag ATP4::tag cytoplasm DAPI
EEF 24h EEFs
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Fig. 22 Spatio-temporal expression of ATP4::mCherry. Fluorescence signal of ATP4::mCherry expressing parasites imaged throughout all stages of the P. berghei lifecycle (a-c). Shown are representative fluorescent and light microscopy images of the respective stages: (a) sexual and asexual blood stages; (b) ookinetes and mosquito midgut stages; (c) 24h, 48h and 69h EEFs after Huh7 infection with sporozoites. Merge of differential interference contrast images (DIC) and cytosolic expressed GFP (green) for parasite identification (1st column);
93 5TH CHAPTER – CRT AND ATP4 merge of ATP4::mCherry signal (red) with Hoechst 33342 nuclear stain (DAPI, blue, 2nd column); ATP4::mCherry signal in greyscale (3rd column). Bar 10 µm.
(a) (b) ATP4-mCherry Sporozoites cytoplasm ATP4::tag ATP4::tag DAPI
60 ± SD 2 m 40
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0
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MGS d14 MGS d15 MGS d16 MGS d17 SGS d18
(c) ATP4::mCherry new localisation
MGS d16 1.0
n 9 8 12 13 14
tes oites no compartment
MGS d17 0.5 f sporoz compartment
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Fig. 23 ATP4::mCherry signal in sporozoites. (a) representative fluorescence images of sporozoites in different developmental stages (MGS: midgut extracted sporozoites, SGS: salivary gland extracted sporozoites). Cytosolically expressed GFP (green) for parasite identification (1st column); merge of ATP4::mCherry signal (red) with Hoechst 33342 nuclear stain (DAPI, blue, 2nd column); ATP4::mCherry signal in greyscale (3rd column). ATP4::mCherry shows a new localisation in MGS d15 and MGS d16 (white arrowhead). Bar 10 µm. (b) integrated ATP4::mCherry live fluorescence density/µm2 analysed for sporozoites with mean (line) and standard deviation (error bars) of respective life stage (day 14 until d18). (c) Ratio of developing MGS and SGS which show a new localisation like indicated with white arrowhead in (a). Number in bars shows total count of sporozoites, n=1.
94 5TH CHAPTER – CRT AND ATP4
5.2.2.2 ATP4 localises in parasite plasma membrane throughout life cycle progression with a new target in midgut sporozoites
For all blood stages and ookinetes, a clear association of the ATP4::mCherry fluorescence signal with the parasite plasma membrane is visible (Fig. 22 a and b). Schizonts display this expression in separated merozoites before the escape from the host cell. Oocysts show a very global fluorescence signal (Fig. 22 b) which cannot be distinguished from mCherry-autofluorescence in wild type parasites without mCherry (Fig. S 4). Sporozoites isolated from mosquito salivary glands show an ATP::mCherry signal in the sporozoite’s plasma membrane. Additionally, targeting to an internal compartment can be observed for some sporozoites extracted from midgut-oocysts predominantly at day 15 and day 16 (Fig. 23 a, white arrowhead). This new localisation is targeted by ATP4::mCherry in the vast majority, but not all day 15 and day 16 midgut sporozoites (Fig. 23 c). Such patterns cannot be seen in autofluorescence of WT sporozoites of equal developmental stage (Fig. S 4). In EEF 24h and 48h p. i. the signal can be detected in the outer membranes, while in EEFs 69h p. i. the signal is mainly identifiable in the plasma membrane of the newly formed merozoites (Fig. 22c).
5.2.2.3 ATP4 shows expression throughout entire life cycle progression
Remarkably, ATP4::mCherry shows a fluorescence signal in all stages, but with different intensities (Fig. 24 a). While in the asexual cycle, a continuous increase can be observed from rings to schizonts, the spatial fluorescence signal remains stable during the differentiation from trophozoites to gametocytes. After the conversion from gametocytes to ookinetes, ATP4::mCherry reaches its strongest expression among all stages. Oocysts show a broad fluorescence intensity variance and a range in negative values due to offsetting with strong autofluorescence in ATP4::mCherry and WT oocysts (compare Fig. S 4 and Fig. S 5). A comparable low signal is seen for sporozoites extracted from salivary glands. As described above, the additional expression in the new localisation observed for midgut sporozoites shows its clearest expression at day 16 (Fig. 23 b). However, sporozoite-emitted fluorescence intensity is in general weaker compared to other life cycle stages. In the Huh7-infection progression, ATP4::mCherry signal is stable but shows an increased variance for the late stages (Fig. 24 a).
95 5TH CHAPTER – CRT AND ATP4
To summarise, ATP4::mCherry shows a global expression with small variations in the expression intensities in the blood stages, and strong variation in oocysts and 69h EEFs. The new localisation observed for midgut sporozoites shows comparable weak intensities.
(a) (b) ATP4::mCherry ATP4 qPCR 1 200 ± SD SP 70 /1) 2 0.1 H m 100 expression
0.01 0 Int. Density / (normalised to relative mRN A -100 0.001
oocysts schizonts EEF 24hEEF 48h ookinetes EEFs 24hEEFs 48hEEFs 69h schizonts ookinetes EEF 72h ringstagesrophozoites sporozoites sporozoites t gametocytes blood stagesgametocytes
Fig. 24 Stage specific expression profiling of ATP4 during life cycle progression. (a) integrated ATP4::mCherry live fluorescence density/µm2 analysed for parasites with mean (line) and standard deviation (error bars) of respective life stage. Oocysts measurements excluded due to strong autofluorescence (b) relative ATP4 mRNA expression measured by qPCR on cDNA converted from RNA of respective life stage (provided by K. Müller). Bars show mean and standard deviation. qPCR was performed in triplets. qPCR has been normalised to the housekeeping gene HSP70/1.
5.2.2.4 ATP4 mRNA expression
To validate the interpretation of ATP4::mCherry fluorescence intensity analysis, additional qPCR was performed on cDNA converted from total RNA extract of the respective life stages (provided by K. Müller) with ATP4 specific qPCR primer combinations. The relative ATP4 mRNA expression is normalised to the housekeeping gene HSP70/1, to which it shows comparable lower expression intensities (Fig. 24 b). Mixed blood stages display the lowest mRNA expression of all stages, directly followed by the most robust expression in schizonts. Gametocytes show a lower expression than schizonts. During ookinete conversion and oocyst development, expression of ATP4 increases. For sporozoites the expression is in a comparable level to blood
96 5TH CHAPTER – CRT AND ATP4 stages. After Huh7 infection, ATP4 mRNA expression shows an increase within three days. Overall, the qPCR expression pattern shows similar trends as the intensity density/µm2 measurements of ATP4::mCherry. Differences in the readout can be seen for a lower expression for ookinetes, and higher expression in 72h EEFs.
5.2.3 Discussion ATP4
ATP4 is a well-studied transporter in asexual P. falciparum blood stages but only poorly described in the remaining stages of the Plasmodium live cycle progression. Drugs inhibiting ATP4’s function describe fast clearance on asexual stages which is important for treating symptomatic infections [162]. Furthermore, ATP4-targeting drugs have been described as powerful gametocidals to interrupt the transmission to mosquitoes [177], indicating the presence of ATP4 and its essential function in this stage. Effects in later stages have been discussed but are overall neglected in the current literature [178]. The ion-homeostatic function of ATP4 in erythrocytes is described in tight dependence and correlation to intra-erythrocytic ion compositions (see introduction). From a cell-physiological perspective, a strong expression variation of a PPM localised sodium pump could be expected during life cycle progression, dependent on the prevailing sodium gradient to either intra or extracellular environments. Therefore, the data of this study can serve to identify a required sodium transport function during life cycle progression, derived from the measured up- and down-regulation of ATP4::mCherry expression.
The qPCR and ATP4::mCherry up- and down regulations in this study progress almost in parallel. However, they differ from described large-scale P. berghei RNA- sequencing data sets: Instead of a peak expression in schizonts, genome wide RNA sequencing shows the strongest ATP4 RNA-expression in ring stages, followed by a continuous expression decrease in schizont and gametocyte stages [148, 149]. The availability of RNA-sequencing data beyond the blood stages is very limited. However, for the P. cynomolgi ortholog a low level of ATP4 expression in EEF-schizonts has been described, while our data suggest an expression comparable to blood stages [178]. Again, a direct translation from RNA-sequencing to qPCR results from different sources is difficult, since our RNA extraction originates from one cell population sample per stage (n=1). Furthermore, the use of large-scale RNA-sequencing and qPCR on
97 5TH CHAPTER – CRT AND ATP4 individual genes are suggested for different experimental approaches which limits a direct translation of the results [179].
The observed ATP4::mCherry expression increase during intra-erythrocytic blood stages suggests a relation to parasite’s increase in size and volume. Consequently, this increase is linked to the production of total membrane proteins, which explains the particular signal increase for membrane-rich schizonts with their internal merozoites. The strong signal in extracellular ookinetes speaks for an upregulation due to environmental changes after the escape from erythrocytes. Sodium concentrations in infected erythrocytes and blood serum differ only slightly [172]. However, after an uptake of blood to the mosquito midgut, changes in sodium concentrations are very likely. A putative upregulation to extrude sodium against an increasing gradient seems reasonable, but remains speculative. To address this, ATP4 dependent ion flux changes in ookinetes could be measured with the use of ATP4 targeting drugs. Unfortunately, this was not possible within the scope of the experimental setup. A complete profile of the sodium concentrations in all intra- and extracellular environments during the parasite’s life cycle progression is to our knowledge not analysed. ATP4::mCherry analysis in oocysts remains limited due to high background and autofluorescence, and is excluded from further interpretation. Nevertheless, the ATP4 mRNA shows a similar expression level like in ookinetes. Sporozoites show the lowest ATP4::mCherry expression, even though they face the most challenging environmental changes. Originating from midgut oocysts they migrate through the mosquitoes hemocoel to the salivary glands, in which they have to remain vital for days up to weeks. Followed by a circulation through the vertebrate host’s bloodstream they finally invade liver cells. The question arises, if sodium extrusion can be down regulated in sporozoites and if yes, why? It was shown that a cytosolic sodium increase in sporozoites induces exocytosis, which is a prerequisite for infection of liver cells [180]. Therefore, it is suggested, that intracellular sodium regulation is interconnected with the sporozoite’s invasion in liver cells [181]. If, and how downregulation of the sodium extrusion is influenced by a lower ATP4 protein- expression in the PPM, remains speculative. Potentially, sporozoites have a weaker global protein expression in general. Remarkably, the selective expression in an unknown localisation of midgut sporozoites, suggests an additional function for ATP4. The fact that this pattern is observed in most, but not all, midgut sporozoites for day 15 and day 16 could imply a regulatory mechanism. It is plausible that ATP4 extrudes or
98 5TH CHAPTER – CRT AND ATP4 imports sodium to the additionally targeted compartment to regulate cytosolic sodium concentrations or even sodium storage. How and if this correlates with emergence from oocysts is unclear. Most importantly, ATP4 targeting drugs show no prophylactic effect on mice infected with sporozoites, indicating either no vital function or no accessibility of the drug to ATP4 in sporozoites [155].
Liver stage localisation and expression intensities are comparable to blood stages. A blood stage-like function in liver stages could, therefore, imply a potential susceptibility against ATP4 directed drugs. The substantial signal variation in 69 hours liver stages, which potentially comes from the three-dimensional size increases and technically inconsistent measurements at this specific stage, limits the interpretation (see technical discussion 5.3). Nevertheless, 48 hours liver stages show a consistent signal intensity. Most important in a physiological context is, if EEFs required sodium extrusion to similar levels as in blood stages, in which a robust erythrocytic influx occurs through the NPP [182]. However, the NPP have so far only been described in infected erythrocytes. Quite in contrast, it was shown that some anti-plasmodial drugs, which are transported via the NPP, show no activity in infected liver cells [183]. In the end, neither NPP influences nor sodium homeostasis in infected liver cells are sufficiently understood to adequately compare the role of ATP4 in blood and liver stages. Nevertheless, the expression in liver stages itself implies a potential function linked to sodium homeostasis.
Altogether, we identify striking P. berghei ATP4::mCherry up-regulations for ookinetes and an expression in sporozoites and EEFs. These new insights not only give hints about a more global function of ATP4, but also suggest a potential activity of ATP4 targeting drugs beyond blood stages.
5.3 Technical discussion
This study describes spatio-temporal full life cycle expression patterns of MTPs as a mean for functional classification of the anti-plasmodial drug-associated MTPs CRT and ATP4. Since many expression analysis techniques are transcriptome based, this approach allows an easily accessible and accurate interpretation of the expression intensities at the protein level without potential underrepresentation of post- translational modifications [184, 185]. The dataset is based on live fluorescence intensity measurements of MTP::mCherry fusion proteins to generate a correlation
99 5TH CHAPTER – CRT AND ATP4 between mCherry intensities and the expression profile of the fused protein. This enables the deduction of quantifiable protein expression along the life cycle progression with simultaneous confirmation of the intracellular localisation. Live cell imaging enables measurements without interfering aspects like fixation of the cells or antibody dilution factors. Nevertheless, influences on the general fluorescence intensity by bleaching or quenching can never be entirely avoided. In particular the ionic environment of the subcellular localisation might have quenching effects on the measured signal (e. g. the digestive vacuole). However, single-cell measurements open the possibility for individual analysis on a cellular basis. The dataset shows, that in most of the stages - in particular blood stages - signal variation is quite low and distinguishable from other developmental stages. An exception are midgut oocysts: since imaging is only possible on oocysts attached to whole midguts, noise and autofluorescence from background and oocysts increase the signal variation in a way which rules out alignments to WT parasites (compare WT signal in Fig. S 4 with MTP::mCherry oocyst signal in Fig. 19 and Fig. 22). An additional factor which limits the inference from mCherry signal to protein expression is the focus depth of three- dimensional objects. Even though the fluorescence signal from deeper/higher foci does not get entirely excluded like in confocal microscopy, signal can get lost by a focus change (indicated in fixed gametocytes in Fig. 18). However, a whole-cell scanning, like in confocal microscopy, would not be feasible for unfixed erythrocytes.
Digital signal processing of the measured intensities per pixel in ImageJ was performed according to the equation in 2.2.2. This approach normalises signal intensity to the measured size while offsetting mean background noise. Measuring only the mean emission might be sufficient for objects with equal size and even signal emission. But in this approach, total protein expression is detected independent of MTP::mCherry’s localisation and most importantly, independent of the size of the parasite or the respective developmental stage. This allows a suitable quantification in which MTP expression is derived from the comparative unit Integrated Density/µm2. The signal integration reflects the sum of intensity values which are integrated over the measured area and gives an intensity output independent of size of the parasite.
To verify this method with a standard expression technique, we carried out qPCR on the respective MTP genes. It has to be noted, that the qPCR data originates from only one biological samples per stage. While this would be insufficient for qPCR-based transcription quantification, it is satisfactory to deal as a verification of the transcription
100 5TH CHAPTER – CRT AND ATP4 trends for MTP::mCherry measurements. As the two techniques reflect either expression at the transcriptome or proteome level, possible delays in mRNA and corresponding protein expression always have to be considered [186, 187]. Additionally, differences in stability of mRNA and proteins, as well as regulatory aspects like mRNA and protein degradation, might have an effect on such a readout. Manipulation of the proteins by mCherry-fusion could lead to a deviated localisation or degradation of the MTP. However, the similar expression progressions for MTP::mCherry and qPCR expression-data for CRT and ATP4 confirm the approach. For example, with CRT, transcript down- and upregulations follow almost identical trends detected in CRT::mCherry signal measurements, with the exception of schizonts. Parallel up- and down-regulations can be observed for ATP4, with the exception of late EEFs (72h). In general, expression intensities show a much higher divergence in qPCR analysis. This describes the limits of the MTP::mCherry analysis.
Together with the technical issues described above, a relative quantification remains limited while overall expression trends, such as up- and down-regulations, can be identified. Therefore, spatio-temporal expression profiling via MTP::mCherry fusion is an appropriate tool to analyse expression trends in life cycle progression, with a simultaneous confirmation of the localisation up to single-cell level.
101 6TH CHAPTER – GENERAL DISCUSSION
6th Chapter: General Discussion 6.1 Findings of this study
This study summarises new functional assignments of MTPs with diverse backgrounds. As shown for Pf3D7_1468600, a basic initial assessment of an MTP- disrupted cell line revealed influences of this protein in asexual blood propagation without being essential. The absence of the protein in P. berghei implies host specific adaptations associated with potential virulence factors in human malaria. In the spatio- temporal expression analysis, the effective drug target P. berghei ATP4 showed an expression beyond blood stages which suggests further testing of spiroindolones in the stages of ATP4 expression. In particular in liver stages, in which the expression reaches almost blood stage-like intensities, prophylactic effects of ATP4 directed drugs could be of high value. Additionally, it shows that the blood stage essential sodium pump maintains its expression across all parasite stages with a deviated localisation in developing sporozoites. The same analysis on P. berghei CRT shows additional expression in liver stages, a stage in which CRT’s localising compartment is not present. A suggested interpretation is a potential early expression of CRT in liver stage merozoites in preparation for the blood stages. If the expression in the liver stages embodies a physiological function of the CRT beyond the digestive vacuole is unclear. And finally, P. berghei FT2 was identified as an apicoplast-resident MTP with a global expression throughout the entire life cycle progression and dual localisation in schizonts and sporozoites. However, an essential contribution of FT2 to the parasite’s development was only identified during sporogony and sporozoite formation. Therefore, deletion of FT2 leads to an arrest of the parasite’s life cycle progression in the mosquito stages. While the physiological role of the defect remains unclear, effects through a folate imbalance on FMT can be excluded.
In summary, the findings reflect functional roles of the studied MTPs in a broad frame. Through the explorative approaches like spatio-temporal expression analysis and gene disruption, potential roles of MTPs during the parasite’s development were suggested. Therefore, the power of experimental genetics allows to study MTPs from initial assessment up to “filling the gaps” of well-studied MTPs.
102 6TH CHAPTER – GENERAL DISCUSSION
6.2 Experimental genetics in Plasmodium parasites
The study applies common methods of gene editing in Plasmodium parasites by targeting MTPs of two species with differing susceptibility to experimental genetics. In P. falciparum the biggest hurdles for experimental genetics are the extreme (A + T)– rich genomic sequence as well as low transfection efficiency and long selection procedures. Nevertheless, constant improvements of established procedures allowed for extensive large-scale gene editing studies with meaningful output in an already early stage of the evolving methods [54]. Together with the implementation of CRIPSR- Cas9 in P. falciparum, editing strategies progressed rapidly [188, 189]. Like shown for the disruption of Pf3D7_1468600, one of the biggest advantages compared to conventional knock out approaches are increased efficiency and shortened selection procedures after transfection. Despite a similar (A + T)-content compared to P. falciparum [190], P. berghei transfections follow advanced selection techniques that make genome editing faster and more feasible compared to P. falciparum [59]. The technical repertoire of P. berghei gene editing strategies allowed me to address functional key aspects, like expression patterns, essentiality and localisations of the MTPs. Additionally, appropriate co-localisation is implementable with GFP expressing organelles. Eventually, an outstanding benefit of the murine malaria model is the easy access of post-erythrocytic stages. Like confirmed by P. berghei FT2 in this study, a genetic screen by Kenthirapalan et al. proposes that the majority of MTPs play a role beyond the blood stages [33], which can suggest extended intervention startegies. Still, one of the major problematic points are genes which are refractory to targeted deletion. After multiple unsuccessful transfection attempts, the genes are often declared as essential (like in [191]). However, the absence of growth is an inconsequent argument for essentiality in asexual blood stages. Therefore, inducible knock downs of the protein production are elegant approaches to study proteins which are essential in asexual blood stages (like in [192]). Exemplified in this study, an inducible knockdown of the unsuccessfully disrupted P. falciparum FT2 is necessary to prove essentiality of the gene in the asexual blood stages.
Despite technical limitations, the improvements made to experimental genetics creates an essential tool to study functional aspects of Plasmodium MTPs on the genome level. While genome editing in P. falciparum was so far reserved for laboratories with advanced know-how, CRISPR/Cas9 now introduces the possibility to
103 6TH CHAPTER – GENERAL DISCUSSION involve experimental genetics into the standard technique repertoire of P. falciparum laboratories.
6.3 The transporter dilemma
The challenges to characterise MTPs are multilayered and the following aspects are of importance when determining the role of an MTP (compare Fig. 25): the transported substrate, the membrane in which the MTP localises, and effects of the MTP on the parasite’s cell physiology, for which changes can be observed after editing the associated MTP gene (derived from [29]). Furthermore, a complete deciphering requires broad experimental approaches, which depend in particular on functional predictions coming from the structural features of the MTP . Amino acid sequence-based predictions can include an identification of functional domains which give hints about a potential substrate. Additionally, a screen for targeting motifs can imply the MTP’s localisation in a specific membrane. For unknown MTPs with a lack of such bioinformatical hints (for instance of the fraction “unknown function” mentioned
2 localisation
3 cell physiological role
4 MTP structure 1 substrates
e. g. involved pathways
external internal
Fig. 25 Four feature-levels which are important for a characterisation of MTPs. (1) The identification of the transported substrate, which can be shown by functional expression in heterologous systems. (2) The membrane in which the MTP localises, shown for instance by experimental genetics. (3) The cell physiological role of for instance the pathways in which the transport is involved. Effects on the cell physiological role can be observed after a manipulation of the gene by experimental genetics. (4) Bioinformatical analysis of structural features which can be used to asses a putative substrate or the localisation by identifying targeting motifs.
104 6TH CHAPTER – GENERAL DISCUSSION in Fig. 2 b), a putative substrate identification can only be derived from the cell physiological context of the experimentally verified subcellular localisation. One example is CRT, for which the expression in the digestive vacuole membrane suggests export related to haemoglobin digestion. Through heterologous expression of MTPs in a suitable system, a group of suspected cargos derived from the subcellular context can be tested for transport, like it was performed for CRT by Bakouh et al. [143]. However, the data of this thesis suggests in three examples, that the subcellular localisation is not always totally consistent during the entire life cycle progression. P. berghei CRT::mCherry, ATP4::mCherry and FT2::mCherry showed a stage-specific deviation of the localising membrane and it is doubtful that the role of an MTP transport is entirely conserved in two different compartments. Therefore, a diverging expression in other membranes, could suggest an extended cell physiological function of the transport, for instance by an involvement of the very same cargo in different pathways. Additionally, promiscuity of Plasmodium MTPs, like recently shown for the P. falciparum hexose transporter HT1 [193], is a phenomenon which can complicate characterisation approaches. Exemplified by FT2, a whole substrate family can potentially be transported, and therefore, multiple substrates of FT2 could be of importance in the apicoplast and the additional target [79].
The latter example summarises a general dilemma between an identification of an MTP transported substrate and the physiological role of the MTPs transport. While often substrate or even transport mechanism are prioritised for a functional characterisation, the pathways in which a cargo is involved and the impact on the parasite’s cell physiology of such pathways, reflect the second layer of the MTP’s function. In the end, a lack of substrate predictions complicates an experimental cargo identification. In return, effects by genetic manipulation of an unknown MTP make an interpretation of the physiological reason for this effect very speculative. In this case, the localisation remains as the only functional hint for undescribed MTPs to cut back the selection of substrates on a few candidates as previously mentioned.
Therefore, an interplay of bioinformatics, substrate identification and experimental genetics is essential to fully characterise MTPs. In particular advancing bioinformatic approaches with more precise structural analysis, for example by profiling hidden Markov model libraries, can lead to a more detailed screen based on domain architecture or taxonomy [194]. A broad revision of putative functions for MTPs with updated prediction tools might be appropriate, since most MTPs are still carrying
105 6TH CHAPTER – GENERAL DISCUSSION annotations from a large-scale MTP analysis performed in 2005 [40] (including the putative aminophospholipid transporter, Pf3D7_1468600). Nevertheless, depending on the complexity of the mechanism, the region of the protein which confers substrate specificity might often be too small to be identified by classical amino acid sequence analysis.
Hence, bioinformatics provides the admission for functional analysis. If this data is lacking, hints derived from the localisation are essential to study the two interdependent aspects, substrate identification and cell physiological role of MTPs.
106 7TH CHAPTER – CONCLUSION AND OUTLOOK
7th Chapter: Conclusion and Outlook
Together, the new findings reveal unknown localisations, expression patterns and involvements in developmental processes of MTPs on Plasmodium’s cell biology. Fluorescently tagged MTPs are a powerful tool for an analysis of expression patterns during life cycle progression with constant confirmation of the intracellular localisation. Assuming a correlation between spatio-temporal expression and the physiological role of the MTP, parasite stages with a putative function of the MTPs were identified. In combination with genetic disruptions or knock out of the associated genes, potential physiological roles were discussed and essential contributions to the parasite’s cell biology highlighted. All studied MTPs, Pf3D7_1468600, ATP4, CRT and FT2 show or have shown a driving contribution to the parasite’s development, which reflects the substantial significance of the protein group.
In following studies, analysis of extensive roles of the selected MTPs in other parasite stages could reveal promising findings. For instance, an assessment of Pf3D7_1468600’s role in sexual blood stages would address a point of immense phospholipid reorganisation and potentially host specific differences in the transmitted parasite stages. In ATP4 and CRT a confirmation of functional roles in the new identified stages of expression would need further studies to support this finding. Additionally, FT2 requires a confirmation of the dual targeting’s localisation and further studies on the substrate and it’s involved pathways which would lead to the observed physiological defects in FT2-deficient mutants.
Other than that, a large majority of Plasmodium’s MTPs lacks studies on functional roles and are potentially hiding momentous information about the impact of the parasites ‘permeome’. Experimental genetics allows for identification of localisations and involvements in physiological processes which need to be complemented with bioinformatical evaluations and extensive substrate studies. Such an inter-methodological approach will lead towards continuing assignments of the role of membrane transport proteins in malaria parasites.
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182. Kirk, K. (2001). Membrane transport in the malaria-infected erythrocyte. Physiol Rev 81, 495-537. 183. Baumeister, S., Wiesner, J., Reichenberg, A., Hintz, M., Bietz, S., Harb, O.S., Roos, D.S., Kordes, M., Friesen, J., Matuschewski, K., et al. (2011). Fosmidomycin uptake into Plasmodium and Babesia-infected erythrocytes is facilitated by parasite-induced new permeability pathways. PLoS One 6, e19334. 184. Rogers, S., Girolami, M., Kolch, W., Waters, K.M., Liu, T., Thrall, B., and Wiley, H.S. (2008). Investigating the correspondence between transcriptomic and proteomic expression profiles using coupled cluster models. Bioinformatics 24, 2894-2900. 185. Hall, N., Karras, M., Raine, J.D., Carlton, J.M., Kooij, T.W., Berriman, M., Florens, L., Janssen, C.S., Pain, A., Christophides, G.K., et al. (2005). A comprehensive survey of the Plasmodium life cycle by genomic, transcriptomic, and proteomic analyses. Science 307, 82-86. 186. Foth, B.J., Zhang, N., Chaal, B.K., Sze, S.K., Preiser, P.R., and Bozdech, Z. (2011). Quantitative time-course profiling of parasite and host cell proteins in the human malaria parasite Plasmodium falciparum. Mol Cell Proteomics 10, M110 006411. 187. Le Roch, K.G., Johnson, J.R., Florens, L., Zhou, Y., Santrosyan, A., Grainger, M., Yan, S.F., Williamson, K.C., Holder, A.A., Carucci, D.J., et al. (2004). Global analysis of transcript and protein levels across the Plasmodium falciparum life cycle. Genome Res 14, 2308-2318. 188. Knuepfer, E., Napiorkowska, M., van Ooij, C., and Holder, A.A. (2017). Generating conditional gene knockouts in Plasmodium - a toolkit to produce stable DiCre recombinase-expressing parasite lines using CRISPR/Cas9. Sci Rep 7, 3881. 189. Xiao, B., Yin, S., Hu, Y., Sun, M., Wei, J., Huang, Z., Wen, Y., Dai, X., Chen, H., Mu, J., et al. (2019). Epigenetic editing by CRISPR/dCas9 in Plasmodium falciparum. Proc Natl Acad Sci U S A 116, 255-260. 190. Hamilton, W.L., Claessens, A., Otto, T.D., Kekre, M., Fairhurst, R.M., Rayner, J.C., and Kwiatkowski, D. (2017). Extreme mutation bias and high AT content in Plasmodium falciparum. Nucleic Acids Res 45, 1889-1901. 191. Dechamps, S., Wengelnik, K., Berry-Sterkers, L., Cerdan, R., Vial, H.J., and Gannoun-Zaki, L. (2010). The Kennedy phospholipid biosynthesis pathways are refractory to genetic disruption in Plasmodium berghei and therefore appear essential in blood stages. Mol Biochem Parasitol 173, 69-80. 192. Charnaud, S.C., Kumarasingha, R., Bullen, H.E., Crabb, B.S., and Gilson, P.R. (2018). Knockdown of the translocon protein EXP2 in Plasmodium falciparum reduces growth and protein export. PLoS One 13, e0204785. 193. Qureshi, A.A., Suades, A., Matsuoka, R., Brock, J., McComas, S.E., Nji, E., Orellana, L., Claesson, M., Delemotte, L., and Drew, D. (2020). The molecular basis for sugar import in malaria parasites. Nature 578, 321-325. 194. Potter, S.C., Luciani, A., Eddy, S.R., Park, Y., Lopez, R., and Finn, R.D. (2018). HMMER web server: 2018 update. Nucleic Acids Res 46, W200-W204.
121 LITERATURE
195. Haussig, J.M., Matuschewski, K., and Kooij, T.W. (2011). Inactivation of a Plasmodium apicoplast protein attenuates formation of liver merozoites. Mol Microbiol 81, 1511-1525. 196. Sato, Y., Hliscs, M., Dunst, J., Goosmann, C., Brinkmann, V., Montagna, G.N., and Matuschewski, K. (2016). Comparative Plasmodium gene overexpression reveals distinct perturbation of sporozoite transmission by profilin. Mol Biol Cell 27, 2234-2244.
122 Acknowledgements
I want to thank Kai Matuschewski for allowing me to perform my research in his working group. He was very encouraging and motivating during my entire PhD and supported me with fruitful discussions and inspiring ideas. Thank you for the trust, your excellent supervision and your engagement for the next generation of researchers. Additionally, I want to thank my second supervisor Alex G. Maier for the warm welcoming in his lab in Canberra. He supported me with exceptional guidance and stimulating feedback while I was learning P. falciparum work during my research stay in Australia. Thank you for this unique experience, your empathy and the introduction into Australia’s outstanding parasitology scene. I especially would like to thank Joachim M. Matz for the remarkable support during my PhD work. He taught me many techniques and mentored me during my work in Germany and Australia. Thank you for your guidance, your patience and the inspiration. And thank you for providing the parasite cell lines for our FT2-project.
I was very fortunate to be a PhD student of the IRTG2290 PhD program, which made this exchange possible and provided excellent research training. The IRTG exposed me to many conferences and allowed me to establish a remarkable network. Thank you to all IRTG members for the stimulating environment and the extraordinary helpfulness. Thank you, Juliane Lippmann and Marylu Grossman for your excellent coordination of the IRTG. Besides, I want to thank Tobias Stauber, who provided great feedback on my work during our TAC meetings. Beyond that, I want to thank the whole Matuschewski and Maier lab for their omnipresent support. Thank you, Katja, Manuel, Alyssa, Theresa, Julie-Anne, Daniela, Merryn and Melanie for your inexhaustible help during my three years of learning. I want to thank Sanketha Kenthirapalan for providing the CRT, and ATP4 parasite cell lines and Julie-Anne Gabelich and Will Hirst for the spell-check of my thesis. Furthermore, I want to thank all IRTG students for the team spirit and friendship we developed already in our very first days.
I want to thank Gemma for her infinite patience during my PhD and her help with my cycle illustration. Finally, I want to thank my family Hildegard, Rémy, Julie and Bernd who made all this possible through their endless trust and constant support.
123 Selbstständigkeitserklärung
Hiermit erkläre ich, die vorliegende Dissertation selbständig angefertigt und keine anderen als die angegebenen Hilfsmittel verwendet zu haben. Wurden Ergebnisse in Kooperationen produziert, ist dies entsprechend angegeben.
Die vorliegende Arbeit wurde an der Humboldt Universität zu Berlin an der Fakultät für Lebenswissenschaften am Institut für Biologie im Institut für molekulare Parasitologie, sowie an der Australian National University in Canberra an der Research School of Biology in der Abteilung Biomedical Science and Biochemistry unter Leitung von Prof. Dr. Kai Matuschewski und Prof. Dr. Alex G. Maier im Rahmen einer Cotutelle- Graduierung durch das Graduiertenprogram IRTG2290 durchgeführt.
Ich erkläre hiermit, dass ich an keiner anderen Universität ein Prüfungsverfahren beantragt bzw. die Dissertation in dieser oder anderer Form bereits anderweitig als Prüfungsarbeit verwendet oder einer anderen Fakultät als Dissertation vorgelegt habe.
François Laurent Frederic Korbmacher
Berlin, den 29. Juli 2020
124 Erweiterte Selbstständigkeitserklärung
Hiermit versichere ich, dass die folgende Publikation
“An apicoplast-resident folate transporter is essential for sporogony of malaria parasites” (eingereicht in Cellular Microbiology) maßgeblich von mir mitverfasst wurde. Mögliche Übereinstimmungen mit Textpassagen aus meiner Dissertation „Towards functional assignment of Plasmodium membrane transport proteins: an experimental genetics study on four diverse proteins“ stellen somit keinen Plagiatsfall dar.
Dies wird bei Bedarf durch die aufgeführten Co-Autoren bestätigt: Benjamin Drepper, Theo Sanderson, Peer Martin, Thomas Stach, Alex Maier, Kai Matuschewski, Joachim M. Matz.
Extended declaration
With this, I guarantee that the following publication
“An apicoplast-resident folate transporter is essential for sporogony of malaria parasites” (submitted in Cellular Microbiology) includes my significant participation. Potential overlaps with passages in the text in my thesis “Towards functional assignment of Plasmodium membrane transport proteins: an experimental genetics study on four diverse proteins“ do not reflect plagiarism.
The listed co-authors can confirm my contribution: Benjamin Drepper, Theo Sanderson, Peer Martin, Thomas Stach, Alex Maier, Kai Matuschewski, Joachim M. Matz.
François Laurent Frederic Korbmacher
Berlin, July 2020
125 Abbreviations % percent °C degree Celsius ~ around µF microfaraday μl microliter μm micrometer μM micromolar bp basepairs cDNA complementary DNA Cas9 CRISPR-associated protein 9 CRISPR clustered regularly interspaced short palindromic repeats C-terminal carboxy-terminal CQ Chloroquine ddH2O distilled water DIC differential interference contrast DI water deionised water DNA deoxyribonucleic acid ds DNA double-strand DNA dNTP deoxyribonucleotide triphosphate E. coli Escherichia coli e.g. exempli gratia EEF exo-erythrocytic form et al. et alia FACS fluorescence activated cell sorting Fig. figure fwd forward g gram g gravity units G guanosine gDNA genomic DNA gRNA guide RNA GFP green fluorescent protein h hours HSP heat shock protein Huh7 Hepato cellular carcinoma cell iRBC infected red blood cell i.p. intraperitoneal i.v. intravenously kb kilobase kg kilogram KO knock out l litre LB Lysogeny Broth M molar m meter mCh mCherry min minutes ml milliliter mm millimeter
126 mRNA messenger RNA MTP membrane transport protein N-terminal amino-terminal n.s. not significant ng nanogram ORF open reading frame P. Plasmodium p p-value PBS phosphate buffered saline p.i. post infection P. berghei Plasmodium berghei P. falciparum Plasmodium falciparum PCR Polymerase chain reaction PPM parasite plasma membrane PV parasitophorous vacuole PVM parasitophorous vacuole membrane RBC red blood cell rev reverse RNA Ribonucleic acid rpm rounds per minute RT room temperature qRT-PCR real-time quantitative PCR s seconds SD standard deviation ssDNA singe-stranded DNA Taq Thermus aquaticus URT untranslated region V Volt v/v volume per volume w/v weight per volume WT wild type
127 Figure Index
Fig. 1 Plasmodium life cycle...... 2 Fig. 2 Membrane Transport Proteins (MTPs) in Plasmodium...... 5 Fig. 3 FT2 is an apicoplast protein and broadly expressed during Plasmodium life cycle progression...... 45 Fig. 4 FT2 shows dual localisation to the apicoplast and to the apical end in segmented schizonts and sporozoites...... 45 Fig. 5 Efficient blood propagation and sexual differentiation in the absence of FT2...... 47 Fig. 6 Ookinete conversion, mosquito midgut colonisation and oocyst development...... 48 Fig. 7 Prominent swelling and cytoplasmic malformations in FT2-deficient oocysts...... 51 Fig. 8 Ectopic expression in ft2– line to rescue sporogonic defects and to assess overexpression of FT2...... 53 Fig. 9 Phenotyping of ectopic expression in ft2– line to rescue sporogonic defects and to assess overexpression of FT2...... 55 Fig. 10 Phenotypic rescue during mosquito infection restores ft2– sporozoite infectivity and reveals redundant functions of FT2 during liver-stage development...... 57 Fig. 11 Experimental genetics approach to interrupt, delete or GFP-tag P. falciparum FT2...... 60 Fig. 12 FMT is an apicoplast protein and exclusively expressed in Plasmodium EEFs...... 64 Fig. 13 Loss of FMT does not lead to a copy of the defect as observed in FT2-deficient parasites...... 65 Fig. 14 Phylogenetic tree of Pf3D7_1468600...... 72 Fig. 15 Experimental genetics approach for CRISPR/Cas9-mediated disruption of Pf3D7_1468600...... 73 Fig. 16 (a) CRISPR/Cas9 transfections with two different gRNA positions on Pf3D7_1468600. .. 74 Fig. 17 Experimental genetics approach for endogenous tagging of CRT...... 82 Fig. 18 CRT::mCherry fluorescence signal co-localises with hemozoin crystals in gametocytes. 83 Fig. 19 Spatio-temporal expression of CRT::mCherry...... 84 Fig. 20 Stage specific expression profiling of CRT during life cycle progression...... 86 Fig. 21 Experimental genetics approach for endogenous tagging of ATP4...... 92 Fig. 22 Spatio-temporal expression of ATP4::mCherry...... 93 Fig. 23 ATP4::mCherry signal in sporozoites...... 94 Fig. 24 Stage specific expression profiling of ATP4 during life cycle progression...... 96 Fig. 25 Four feature-levels which are important for a characterisation of MTPs...... 104
128 Table index
Tab. 1 PCR component-mix for high fidelity (HF) and Taq-polymerase in P. berghei (P.b.) and P. falciparum (P.f.) ...... 15 Tab. 2 Time and temperature settings for P. berghei (P.b.) and P. falciparum (P.f.) polymerase chain reaction for high fidelity and Taq-polymerase...... 16 Tab. 3 Set up of restriction digest for 50 µl...... 17 Tab. 4 Composition of qPCR master mix ...... 21 Tab. 5 qPCR protocol with time and temperature steps ...... 21 Tab. 6 Parameter settings for P. falciparum electroporation in BioRad Genepulser Xcell ...... 34
Supplementary figure index
Fig. S 1 Diagnostic PCR to test for wild type (WT) or integration specific (INT) PCR products as indicated in Fig. 15 b...... 130 Fig. S 2 Detailed vector map of pDC2-cam-coCa9-U6 plasmid with homologous regions of Pf3D7_1468600...... 131 Fig. S 3 Endogenous tagging of FT2 in P. berghei...... 132 Fig. S 4 WT Bergreen autofluorescence in mCherry channel imaged throughout all stages of the P. berghei lifecycle...... 133 Fig. S 5 ATP4::mCherry fluorescence signal for oocyst stages...... 134 Fig. S 6 Detailed vector map of pCC1 and gGlux6 plasmid tailored to approach P. falciparum FT2...... 135 Fig. S 7 Assessment of fmt– growth compared to WT parasites...... 136
Supplementary table index
Tab. S 1 Oligonucleotides used for P. berghei...... 137 Tab. S 2 Oligonucleotides used for P. falciparum...... 138
129 Appendix
Pf3D7_1468600 Pf3D7_1468600 disrupted 1 disrupted 2 wild type
kb 5‘ WT 3‘ WT 5‘ INT 3‘ INT 5‘ WT 3‘ WT 5‘ INT 3‘ INT 5‘ WT 3‘ WT 5‘ INT 3‘ INT 3.0 2.0 1.5
1.0
0.5
Fig. S 1 Diagnostic PCR to test for wild type (WT) or integration specific (INT) PCR products as indicated in Fig. 15 b. Pf3D7_1468600 disrupted 1 cell line indicates correct integration and absence of wild type locus. Pf3D7_1468600 disrupted 2 cell line shows unspecific bands in 5’INT, correct integration in 3’INT and absence of wild type locus. Wild type control confirms 5’ and 3’WT products and no product for integration specific primer combinations.
130
Fig. S 2 Detailed vector map of pDC2-cam-coCa9-U6 plasmid with homologous regions of Pf3D7_1468600. The vector map shows unique REase sites and the BbsI site which is used for gRNA ligation. Purple sites are annealing sites for sequencing primers or primers for diagnostic PCR (genotyping Primers). Pf3D7_1468600 homologous regions flank hDHFR drug cassette under 5’PcDT promotor. Cas9 codon optimised for P. falciparum (coCas9) expressed under calmodulin promotor (CAM). Modified gRNA expression, harbouring targeted gRNA sequence, under U6 promotor (P. falciparum U6 snRNA). Ampicillin resistance cassette (AmpR) for drug selection during molecular cloning. Note that BbsI is not unique and used for ligation of tailored gRNA sequence. The total size of the modified plasmid: 12.197 bp.
131 (a) WT
P1 P2 FT2
FT2 (CT) tag api-GFP cassette drug cassette
5‘INT 3‘INT
P1 P3 P4 P2 FT2 tag api-GFP cassette drug cassette FT2 (CT)
(b) (c) INT ring stage trophpzoite schizont
WT 5‘ 3‘ - 2.0 kb ft2-tag FT2::tag
- 0.5 kb - 2.0 kb DNA FT2::tag ft2-tag + apiGFP
- 0.5 kb DIC
Fig. S 3 Endogenous tagging of FT2 in P. berghei. (a) Strategy for the generation of transgenic parasites that express the endogenous FT2 fused to mCherry-3xMyc (tag, red) by single homologous integration. Recombinant parasites express the drug-selectable hDHFR-yFcu cassette (blue) and a cassette driving expression of cytoplasmic (not shown) or apicoplast-targeted GFP (api-GFP, green, shown), resulting in the ft2-tag and ft2-tagapi- GFP parasite lines. Wild type-specific (WT) and integration-specific (INT) primer combinations are indicated by arrows and expected fragments by dotted lines. Shown are the WT locus (top), the transfection vector (middle) and the recombined locus (bottom). Cloning and generation of isogenic FT2 ::tag cell line was done by J. M. Matz. (b) Diagnostic PCRs of the mixed parasite populations post-transfection, using the primer combinations indicated in (a). Results are shown for the ft2-tag (top) and the ft2-tagapi-GFP parasite lines (bottom). Arrowheads show expected product size (details in Tab. S 1). (c) Live fluorescence microscopy reveals that FT2 localises to a dividing organelle during blood stage development. Shown are the fluorescent signal of tagged FT2 (red, 1st row), a merge with Hoechst 33342 nuclear stain (DAPI, blue, 2nd row), and transmitted light images (3rd row). Imaging performed by J. M. Matz. Bar, 5 µm
132 (a) DIC mCherry DIC mCherry (b) mCherry mCherry cytoplasm DAPI cytoplasm DAPI
ookinete
ring stage
trophpzoite
oocyst d10 blood stages
schizont
oocyst d14 ookinetes and mosquito stages
oocyst d17
gametocyte
(c) DIC mCherry (d) 100 WT autoflourescence mCherry cytoplasm DAPI 80 ± SD 2
m 60
40
20 Int. Density /
sporozoite
0
Schizont Ookinete Ringstage Sporozoite Trophozoite Gametocyte
EEF 24h (e) (f) WT autoflourescence WT autoflourescence 2000 2000 sporozoitres 1500 1500 ± SD ± SD 2 2 m m
EEF 48h 1000 1000
500 500 Int. Density / Int. Density /
0 0
EEF 69h
scale bar: 10 µm Liver 24hLiver 48h Liver 69h Oocysts d10Oocysts d14Oocysts d17
Fig. S 4 WT Bergreen autofluorescence in mCherry channel imaged throughout all stages of the P. berghei lifecycle. (a-c) Merge of differential interference contrast images (DIC) and cytosolically expressed GFP (green) for parasite identification (1st column); merge of WT-mCherry signal (red) with Hoechst 33342 nuclear stain (blue, 2nd column); WT-mCherry signal in greyscale (3rd column). Imaging in equal mCherry exposure times as for ATP4::mCherry and CRT::mCherry. Note high
133 autofluorescence in oocysts (f). (d-f) integrated wild type live autofluorescence intensity/µm2 analysed for respective parasite stage. The mean was defined as autofluorescence and subtracted from signals measured in CRT::mCherry and ATP4::mCherry measurements.
ATP4-mCherry 1000
500 ± SD 2
m 0
-500
-1000 Int. Density /
-1500
OocystsOocysts d10 Oocysts d14 d17
Fig. S 5 ATP4::mCherry fluorescence signal for oocyst stages. Integrated ATP4::mCherry live fluorescence intensity/µm2 analysed for oocysts. WT autofluorescence of respective stage was subtracted. Plot shows mean plus standard deviation.
134 (a)
(b)
Fig. S 6 Detailed vector map of pCC1 and gGlux6 plasmid tailored to approach P. falciparum FT2. Map shows all components of the plasmid plus used REase sites which were used for molecular cloning. (a) pCC1 plasmid with drug cassette (hDHFR) under CAM 5’ promotor flanked by homologous regions (HR1 FT2 and HR2 FT2). Molecular cloning was performed via Gibson assembly. pCC1 plasmids harbour additionally a CD cassette under hsp85 5’ promoter for negative selection. Purple notes show the annealing sites for sequencing primers. pGEM3Z backbone carries Ampicillin resistance cassette for drug selection during molecular cloning. MCS indicates
135 REase site-rich cloning sites. Total size of the modified plasmid: 8,555 bp. (b) pGlux6 plasmid with P. falciparum FT2 expressed under CRT 5’ promotor with a C-terminal GFP tag. REase site used for molecular cloning: KpnI and XhoI. Plasmid carries a modified PAC cassette under CAM 5’ promotor for drug selection against puromycin. Backbone carries Ampicillin resistance cassette for drug selection during molecular cloning. Total size of the modified plasmid: 9.096 bp.
(a) (b) (c) 101 n.s. 50 100 100 40 80 10-1 30 60 10-2 20 40 10-3 Parasitemia (% ) WT 10 20 10-4 fmt- 0 Ookinete conversion (% ) 0 3 4 5 6 7 WT fmt– WT fmt– Gametocyte conversion rate (% ) Days post infection
(d) (e)
t 1000 2000 2 800 1500
600 1000 400
d17 are a in µm 500 200
Oocysts / infected midgu 0 0 WT fmt– WT fmt–
Fig. S 7 Assessment of fmt– growth compared to WT parasites. (a) Intravital competition assay. Equal numbers of mCherry-fluorescent Berred WT and GFP- fluorescent fmt–blood stage parasites were co-injected intravenously into mice and peripheral blood was analysed daily by flow cytometry [60]. n.s., non-significant; two-way ANOVA to test for the two variables time and genotype; n=3. performed by J. M. Matz. (b) Sexual differentiation in the absence of FMT is not affected. 107 WT or fmt– blood stage parasites were injected intravenously into mice and peripheral blood was analysed three days later for gametocyte conversion; n=1. (c) Percentage of fully matured ookinetes among in vitro cultivated P28-positive parasites appears normal in absence of FMT; n=1. (d) Mosquito midgut colonisation in the absence of FMT comparable to WT parasites. Shown are oocyst numbers of infected midguts and the mean infection intensity (lines) on day ten after the blood meal; (n=1). (e) Area occupied by oocyst was imaged live 17 days after the blood meal from one blood meal. Shown are individual data points as well as mean values. Swelling in late stages does not occur on 2D Area level.
136 Tab. S 1 Oligonucleotides used for P. berghei. Underlined sequence: restriction enzyme, WT: length of PCR product (bp) on wild type genomic DNA, INT: length of PCR product (bp) on mutant’s genomic DNA, TV: transfection vector, GT: diagnostic genotyping PCR, SK: Sanketha Kenthirapalan, JMM: Joachim M. Matz.
Primer sequence WT INT Use Target Reference CT-PbATP4-F-SacII aaaccgcggcttaatttttgggacccacgag TV CT-PbATP4 SK 592 CT-PbATP4-R- aaagacatatgtccattttttattgacatatattttcttctgtatataac CT-PbATP4 TV SK PshAI 3’- PbATP4-F KpnI ataggtaccttcatccactttatgttgaaatgag TV 3’-PbATP4 SK 515 3’- PbATP4-R-AvrII aatcctaggtgtgcaaatgtatgtgtgtaatcagg TV 3’-PbATP4 SK CT-PbCRT-F-SacII aaaccgcggaaattcgaaatattttccccttatttttac TV CT-PbCRT SK 529 CT-PbCRT-R-PshAI aaagacatatgtcctgcccttgatgtttctatagaag TV CT-PbCRT SK 3’- PbCRT-F KpnI ataggtacctgccacataaagttataaagaagtatg TV 3’-PbCRT SK 567 3’- PbCRT-R-AvrII atacctaggaggcaaataaggatataagacaaaag TV 3’-PbCRT SK CT-PbFT2-F-EcoRI aattgtgaattcttggaatgcgaagaaaaccg TV CT PbFT2 JMM 1079 CT-PbFT2-R-HpaI ttttttgttaactttctgtatttttttattttttaaactttcaatttg TV CT PbFT2 JMM 5’-PbFT2-F-SacII aataatccgcggccaacatttccatctcgcg TV 5’ PbFT2 JMM 1069 5’-PbFT2-R-PvuII tttaatcagctggtatgtaactattcgtatgtgcg TV 5’ PbFT2 JMM 3’-PbFT2-F-XhoI aataatctcgagaaaaaatacagaaatgagcggc 682 TV 3’ PbFT2 JMM 3’-PbFT2-R-KpnI aataatggtacctaaacggtttggatgaagcg TV 3’ PbFT2 JMM 5’-CPN20-F-PvuII gctaagccagctgaaaaaactaacaatatttatttgacgc 1361 TV 5’ PbCPN20 JMM NT-CPN20-R-PshAI actcacagacatatgtcctctaattgttctattatctagc TV NT PbCPN20 JMM 5’-CPN20-F-SacII attaatccgcgggtcctattctctcaacgatgg TV 5’ PbCPN20 JMM 1085 5’-CPN20-R-EcoRI aaaagtgaattcgcataaatatttattgtatattcatatagtttag TV 5’ PbCPN20 JMM 5‘-PbFMT-F-SacII aaattaccgcgggtctatgaaggttttgaaaatcgg TV 5‘ PbFMT JMM 823 5’-PbFMT-R-EcoRI ttttatgaattctatttgcctgtgcaacattcc TV 5‘ PbFMT JMM 3‘-PbFMT-F-XhoI aataaactcgagaaaataagcaaattgtggatgcc TV 3‘ PbFMT JMM 868 3‘-PbFMT-R-KpnI aataatggtaccgaaaataagtttagcaaaagttaatgc TV 3‘ PbFMT JMM CT-FMT-F-EcoRI tttaaagaattcaaaatgttatcacaaaaagccccc TV CT PbFMT JMM 1169 CT-FMT-R-HpaI tttacagttaacatataatataactccacgattaatgctattaac TV CT PbFMT JMM 5’-PbATP4-F tgacaaaaagatggtggttatatgg GT 5’ PbATP4 SK 803 5’-PbATP4-R ctgtattttggccattttcaacg GT 5’ PbATP4 SK 3’-PbATP4-F tgacaaaaagatggtggttatatgg 948 GT 3’ PbATP4 SK 1670 3’-PbATP4-R ataattttcttttcagattgatcgc 1456 GT 3’ PbATP4 SK 5’-PbCRT-F attgtttgttttataggagtcgacc GT 5’ PbCRT SK 929 5’-PbCRT-R cattcgagcttaattgtttgacc GT 5’ PbCRT SK 3’-PbCRT-F aggaccagccataacaatagc 704 GT 3’ PbCRT SK 1198 3’-PbCRT-R gtcttctaaacaacgagcatg 1228 GT 3’ PbCRT SK 5’-PbFT2-F gtttcttttacctgtttcttttaccc 1428/1807 GT 5’ PbFT2 JMM 1376 5’-PbFT2-R aatttatcaaaggcaattcttctcc GT 5’ PbFT2 JMM 3’-PbFT2-F ggctccctcacaaaaaaggg GT 3’ PbFT2 JMM 926 3’-PbFT2-R tgcccgaaaaatagaagtccc 1377 GT 3’ PbFT2 JMM CT-PbFT2-F gattaatacatttagcatctttggc 1318 GT CT PbFT2 JMM 1342 CT-PbFT2-R attggagcattttttttgcgc 2011 GT CT PbFT2 JMM 5’-PbFMT-F tcaatgacagaaataaaaggtagc GT 5‘ PbFMT JMM 1188 5’-PbFMT-R tgtcctcccaattattcacccc GT 5‘ PbFMT JMM 3’-PbFMT-F acaagtgttttgatggcattcc GT 3‘ PbFMT JMM 1020 3’-PbFMT-R cttcaattggatcaccttcacc GT 3‘ PbFMT JMM CT-PbFMT-F aaaagaaagatgatattgtagatatgc 2196 GT CT PbFMT JMM SIL6F gacagcgcatatgatggatg 1315 1804 GT PbSIL6 [59] SIL6R tacgaatacgcaatttctcaaac 1247 GT PbSIL6 [59] 5’DHFRrev atgaaataccgctccatttttcc GT 5’ PbDHFR-TS [59] 5’HSP70rev caatttgttgtacataaaataggcag GT 5’ PbHSP70 [59] 3’DHFS-F gcttttcgctatatcgctatc GT 3’ PbDHFS-FPGS [52] 5’-PbHSP70-3-R catgataacccattactaatatgtgg GT 5’ PbHSP70-3 JMM mCherryRev ccctccatgtgaaccttgaag GT mCherry [195] M13R caggaaacagctatgaccatg GT M13 [195] PbHsp70-F gcaaagccaaacttaccagagtc qPCR PbHSP70 [196] 125 PbHsp70-R gttctattaccttggtcgtttgc qPCR PbHSP70 [196] PbATP4-F agcaaagcgtgattccaaacc qPCR PbATP4 FK 277 PbATP4-R tcagtttgggttctttcgacc qPCR PbATP4 FK PbCRT-F attgttgagaactgtggtgttgg qPCR PbCRT FK 220 PbCRT-R gttttcttacagcatcgcccg qPCR PbCRT FK
137 Tab. S 2 Oligonucleotides used for P. falciparum. CAPITALISED sequence: Gibson homologous regions, WT: length of PCR product (bp) on wild type genomic DNA, INT: length of PCR product (bp) on mutant’s genomic DNA TV: transfection vector, GT: diagnostic genotyping PCR, FK: Francois Korbmacher.
Primer sequence WT INT Use Target Reference NT-PfFT2-F-XhoI tatactcgagatgatagaaaagtctaacaatcc TV NT-PfFT2 FK 1385 CT-PfFT2-R-Kpni attaccatggtcccttggatgtttcc TV CT-PfFT2 FK TATATATCCAATGGCCCCTTTCCGCgggag NT-PfFT2-F-GIB TV NT-PfFT2 FK ccttccatttaaagg 582 ATGCTTAAGACAGATCTTCGGAccacagttgt NT-PfFT2-R-GIB TV NT-PfFT2 FK acaaaaggatg ATAAGAACATATTTATTAAATCTAgaattcgatt CT-PfFT2-F-GIB TV CT-PfFT2 FK cagaccatcgtttatggg 554 ATACCGCATCAGGCGCCAGCcgatgtgctcctc CT-PfFT2-R-GIB TV CT-PfFT2 FK gt TATGGGAATTTCCTTATAGGGCCCgggagc NT-PfFT2-F-GIB TV NT-PfFT2 FK cttccatttaaagg 387 TAGCTAAGCATGCggttttaatataaaagggatatat NT-PfFT2-R-GIB TV NT-PfFT2 FK ggatac CT-PfFT2-F-GIB TAGAGGTACCGAgattcagaccatcgtttatggg TV CT-PfFT2 FK CCCCGAAAAGTGCCACCTGACGTCggtggc 359 CT-PfFT2-R-GIB TV CT-PfFT2 FK aaaaacactagc GGAATTTCCTTATAGGGCCggaaaacgtacaa NT-Pf3D7_1468600-F-GIB TV NT-Pf3D7_1468600 FK acgacttacac 489 NT-Pf3D7_1468600-R-GIB TAAGCATGCcctttcattcgatcaaaatggc TV NT-Pf3D7_1468600 FK CT-Pf3D7_1468600-F-GIB CTCGAATTCgggatgaatgtaggcttattcaaag TV CT-Pf3D7_1468600 FK AAAAGTGCCACCTGACGTggtttgaaagaagag 686 CT-Pf3D7_1468600-R-GIB TV CT-Pf3D7_1468600 FK agcactatatg NT-Pf3D7_1468600-F gaagagagaacatatcaaacag 625 GT NT-Pf3D7_1468600 FK 523 NT-Pf3D7_1468600-R gtaacctcactagagtttaacaac GT NT-Pf3D7_1468600 FK CT-Pf3D7_1468600-F gtgatagttgtagttatagtagcg GT CT-Pf3D7_1468600 FK 1452 CT-Pf3D7_1468600-R cttttgtaatgtgtccacaac 1202 GT CT-Pf3D7_1468600 FK 5’-PcDT-R cacatgcttagtacacatgc GT 5’-PcFT FK 3’-HRP2-F ccgcagagaaatctagagg GT 3’-PfHRP2 FK NT-PfFT2-F cttagaagaaataattggggagc 1572 GT NT-PfFT2 FK 658 NT-PfFT2-R gcaactgataaactacctattgc GT NT-PfFT2 FK CT-PfFT2-F gccacttccatatttccg GT CT-PfFT2 FK 759 CT-PfFT2-R cattatttatcccttggatgtttcc 860 GT CT-PfFT2 FK
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