Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of Trypanosoma brucei

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Thesis

Reference

Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of Trypanosoma brucei

AMBUEHL, Chiara

Abstract

Human African trypanosomiasis caused by the infection of Trypanosoma brucei (T. brucei) is a fatal disease if untreated. Today treatments are either very toxic, very complicated to administer or limited to the first phase of the infection, thus new potent and safe drugs are needed. The aim of the thesis was to investigate the AMP-activated protein kinase (AMPK) in T. brucei. AMPK is a heterotrimeric complex known for its role in maintaining energy homeostasis in the cell by detecting the ratio AMP:ATP. To this end the corresponding subunits were identified by sequence alignment and confirmed by a modified bacterial adenylate two-hybrid system : catalytic subunits α (TbAMPKα1: Tb927.3.4560, TbAMPKα2:

Tb927.10.5310) and regulatory subunits β (Tb927.8.2450) and γ (Tb927.10.3700). The complex was expressed using a polycistronic expression system and the purified complex was biochemically characterized. Finally, the importance of the catalytic subunits were confirmed in vivo on designed knock-out and knock-down strains.

AMBUEHL, Chiara. Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of Trypanosoma brucei. Thèse de doctorat : Univ.

Genève, 2018, no. Sc. 5239

DOI : 10.13097/archive-ouverte/unige:110627 URN : urn:nbn:ch:unige-1106277

Available at:

http://archive-ouverte.unige.ch/unige:110627

Disclaimer: layout of this document may differ from the published version.

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Section des Sciences Pharmaceutiques Professeur Leonardo Scapozza Laboratoire de Biochimie Pharmaceutique Docteur Remo Perozzo

Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of

Trypanosoma brucei

THÈSE

présentée à la Faculté des sciences de l’Université de Genève

pour obtenir le grade de Docteur ès sciences, mention sciences pharmaceutiques

par

Chiara Ambühl de

Uznach (SG)

Thèse Nº 5239

GENÈVE

Atelier d’impression ReproMail 2018

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A Léa

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Acknowledgments

At the end of this journey which was my doctoral thesis, I cannot help but thank all those who have contributed in one way or another to this adventure.

First of all, I would like to thank my two thesis co-directors for giving me the opportunity to do my thesis in the very multi-faceted and dynamic Pharmaceutical Biochemistry group.

I would like to thank Professor Leonardo Scapozza for all the advice and discussions during the meetings and presentations, as well as for all the interesting conversations not work-related. Thanks to Dr Remo Perozzo, my direct supervisor, for having guided me on this long road while allowing me great freedom in the conduct of my experiments. Thank you for your great patience and for the great knowledge you were able to share with me, especially during our meetings, which always started with a "you have 5 minutes...?" and have always lasted much longer. All these recommendations have allowed me to progress, even and especially when the results were not what we had hoped for.

I would then like to thank the external members of my thesis committee, Professor Pascal Mäser, Dr Emerson Ferreira Queiroz and Dr Brooke Morriswood, for taking the time to read and comment my work.

The joyful and peaceful environment within the laboratory has been a very important energy resource during these years and is the result of the presence of many great people. Thank you all for the interesting discussions scientific and not, thank you for all the advice scientific and not and for the good moments spent together in the laboratory or outside.

I would first of all like to thank Patricia Graven for integrating me into the world of trypanosomes and for the good times spent together these past years. I would particularly like to thank Magali Zeisser-Labouèbe, my office colleague, for the good friendship she has shown me, for the moments shared during the working days and beyond, and for the chatting during coffee breaks or the "pineapple-basil ice cream"

getaways.

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The Aurelie's, Aurélie Baguet and Aurélie Gouiller, thank you for the good memories of time spent together at noon, in the laboratory and outside.

A special memory will remain for the now historic "TROPICAL LAB", in which we suffered endless hours in summer and winter but thanks to all of its members, there were also many laughs and remains today a good memory. Hesham Ismail, thank you for the good discussions and laughter, and for the difficult challenges you gave me to translate the French songs that were played on the radio in the cell culture lab. Thanks also for keeping my trypanosomes alive when I was not in the lab.

Verena Santer, our Italian and German chats have always put me in a good mood and I already miss them. Elinam Gayi for the happiness and energy that always radiates in your presence. Francesca Tessaro and Amparo Garcia for the nice talks, for your advices and for the laugh together.

Andreja Vujicic Zagar and Oscar Vadas and Piter, thanks for all the valuable advice and good discussions and laughs in the lab and outside. Béatrice Kaufmann, Laurence Neff and Olivier Dorchies for all your precious advices and your availability. Karl Perron and Verena Ducret for the wonderful atmosphere during the practical bacteriology work for Bachelor students. Sarah Gay, my master's student, for her professionalism and excellent work done. Charlotte Petit for her good friendship and the beautiful evenings together at the theatre discovered in the basement of your old building.

I would also like to thank all the other past and present members of the various Pharmacy groups, FABIP, FAPER, FABIO, FATHO, FAMCO for the wonderful time spent together.

Today, I would not be here to write these thanks if I had not the support of all my friends. The people from Geneva that I met during my university studies and outside (Drine, Mel, Youyou, Fanny, Marion, Annick, Vania, Jerem, Snouss, Cucu, Martin, Jule, Ludo, Yann, Patrick, ...) and the people from Ticino that are now scattered all over Switzerland (Gioia, Nora, Sheila, Marco, ...). Thank you for everything, thank you for your moral support in the most difficult moments and for the friendship you know how to show me.

In the end, I would like to thank my parents, Liz and Ferruccio, for giving me the chance to study and make my own life choices. Thanks for all the support during those years

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away from home and even more thanks for all the help this past year, without you, the writing would have been even more difficult. Alessandro and Floriane, thank you very much. Thanks for all the advices and thanks for the patience and technical support you gave me every time I called and said "I have a big and ‘urgent problem’!”.

Thanks also to the great and big "Belle-Famille" who welcomed me since more than 10 years with open arms.

Finally, I would like to thank Anthony and Léa. Anthony thank you for the last 10 years of life together and thank you for all your love and support (especially in these last months). Léa, you certainly didn't help me finish the thesis, on the contrary, but your energy and tenderness is a gift every day.

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Abbreviations

AAT African Animal Trypanosomiasis

ADP Adenosine 5'-diphosphate

AMP Adenosine 5'-monophosphate

ATP Adenosine 5'-triphosphate blastR Blasticidin resistance cassette

bp Base pair

BSF Bloodstream form

CAMKK2 Calmodulin-dependent kinase kinase 2 cAMP Cyclic adenosine monophosphate

CATT Card agglutination test for trypanosomiasis CBP Calmodulin binding peptide

cDNA Complementary DNA

CNS Central nervous system

CSF Cerebrospinal fluid

DNA Deoxyribonucleic acid

DNDi Drug for Neglected Diseases Initiative E. coli Escherichia coli

EDTA Ethylenediamine tetraacetic acid EP/GEET Procyclines on the surface of PCF

EtOH Ethanol

FCS Fetal Calf Serum

gDNA genomic DNA

h Hours

HAT Human African Trypanosomiasis HMI-9 Hirumi’s modified Iscove’s medium 9 HPLC High performance liquid chromatography hygroR Hygromycine resistance cassette

IPTG Isopropyl-β-D-thiogalactopyranoside

K_M Michaelis constant

LKB1 Tumor suppressor kinase LKB1 MCS Multiple Clooning Site

min Minutes

mRNA Messenger RNA

NGO Non-governmental organizations NPO Non-profit Organizations

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Nysm T. b. brucei New York single marker cell line PAGE Polyacrylamide gel electrophoresis

PCF Procyclic form

phleoR Phleomycin resistance cassette PMSF Phenylmethylsulfonyl fluoride PuroR puromycine resistance cassette

RBS Ribosome Binding Site

RDT Rapid Diagnostic test

RNA Ribonucleic acid

RNAi RNA interference

RT Room temperature

s Seconds

SD Standard deviation

SDS Sodium dodecyl sulfate

SEC-MALS Size exclusion chromatography coupled to multi angle light scattering SRA Serum resistance-associated

SSF StrepII-StrepII-FLAG tag SSH StrepII-StrepII-3xHA tag T. b. brucei Trypanosoma brucei brucei T. b. gambiense Trypanosoma brucei gambiense T. b. rhodesiense Trypanosoma brucei rhodesiense T. brucei Trypanosoma brucei

TAK1 Growth factor-β-activated kinase 1 TAP Tandem affinity purification TbAK T. brucei adenosine kinase

TbBip Homologous to immunoglobulin heavy chain binding protein TetR Tetracycline repressor

TEV Tobacco etch virus

TLF Trypanosoma lytic factor

TOR Target of rapamycin

TRIS Tris(hydroxymethyl)aminomethane

UTR Untranslated region

VSG Variant surface glycoprotein WHO World Health Organization

ZMP AICAR monophosphate

β-CBM β-Carbohydrate binding module or glycogen binding domain

λPP Lambda protein phosphatase

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Résumé de la thèse

La trypanosomiase humaine africaine (THA), également appelée maladie du sommeil, est une maladie causée par l'infection de Trypanosoma brucei (T. brucei), un parasite unicellulaire appartenant à l'ordre des Kinetoplastida vivant en Afrique subsaharienne et transmise par la mouche tsétsé du genre Glossina. Deux sous-espèces de T.

brucei sont responsables de la maladie, T. b. gambiense responsable de la forme chronique représentant 97% des infections et T. b. rhodesiense responsable de la forme aiguë représentant les 3% restants.

Aujourd’hui, les traitements contre cette maladie sont soit très toxiques, soit très compliqués à administrer ou limités à la première phase de l’infection en raison des difficultés à franchir la barrière hémato-encéphalique, étape nécessaire au traitement de la deuxième phase de l’infection. Pour ces raisons, la découverte de nouvelles cibles thérapeutiques et/ou de nouveaux médicaments capables d’éliminer T. brucei à tous les stades d’infections est primordiale pour éradiquer la maladie du sommeil. Ce travail de thèse a été entreprise après la découverte d’un possible lien entre T.

brucei adénosine kinase (TbAK) et la cascade de la protéine kinase activée par l'AMP (AMPK). L'AMPK est un complexe hétérotrimérique connu pour son rôle dans le maintien de l'homéostasie énergétique dans la cellule en détectant le rapport AMP:ATP. Il est composé d'une sous-unité catalytique α devant être phosphorylée afin d’être activée et de deux sous-unités régulatrices β et γ.

Sur la base de cette observation nous avons décidé d'étudier T. brucei AMPK (TbAMPK), complexe jouant un rôle primordial dans la survie de la cellule et très peu caractérisé chez T. brucei, particulièrement dans la forme sanguine du parasite (Blood stream form, BSF).

Au cours de cette étude, nous avons confirmé l’identité des sous-unités régulatrices TbAMPKβ (Tb927.8.2450) et TbAMPKγ (Tb927.10.3700), précédemment décrites dans la forme procyclique (forme caractéristique du parasite dans le vecteur) également dans les parasites BSF. D’autre part, nous avons démontré que les deux candidats de la sous-unité catalytique α (α1: Tb927.3.4560, α2: Tb927.10.5310), identifiés par alignement de séquence entre les sous-unités α humaines et le génome du parasite, formaient bien un complexe avec TbAMPKβ et TbAMPKγ. Ceci a été

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possible en modifiant la technique double hybride de l’adenylate cyclase (BACTH system) à la détection d’une interaction entre deux partenaires dans un complex multi- protéique. Ce nouvel outil nous a permis de démontrer que les deux candidats de la sous-unité catalytique formaient bien un complexe avec les autres sous-unités.

Après avoir identifié les sous-unités, le complexe a été exprimé grâce à l’utilisation d’un système d’expression polycistronique intégrant les trois sous-unités dans le même plasmide sous le contrôle d’un promoteur unique. Cette méthode a permis de produire avec succès le complexe TbAMPKγβα2 dans des bactéries. Le meilleur rendement a été obtenu après induction par 1 mM d’IPTG de la production des protéines à 12°C pendant 72 h. Grâce à la présence d’un His6-tag en N-terminus de la sous-unité catalytique, le complexe a pu être isolé en utilisant une chromatographie d’affinité au Nickel.

Afin d’améliorer la pureté du complexe produit, une chromatographie d’échange d’ion et une chromatographie d’exclusion de taille (SEC) ont été appliquées par la suite nous permettant de produire 0.16 mg de complexe de protéine par g de masse de bactéries initiale. L’analyse par SEC-MALS (Multi angle light scattering) a permis de confirmer que le poids moléculaire absolu du complexe purifié était de 160 kDa comme attendu.

Par la suite, l’évaluation de l’état de phosphorylation nous a permis de mettre en évidence que le complexe exprimé dans la bactérie était déjà phosphorylé au niveau du site crucial de phosphorylation de la sous-unité catalytique, Thr172. De plus, une incubation supplémentaire avec une AMPK kinase n’a pas augmenté le niveau de phosphorylation. De même, l’incubation du complexe purifié ou la production du complexe en présence d’une Lambda phosphatase (λPP) n’a pas permis de réduire le signal de phosphorylation détecté au niveau de la sous-unité catalytique.

L’utilisation d’un essai de bioluminescence permettant de déterminer le niveau d’ADP produit par la réaction de la kinase en présence d’un substrat spécifique (SAMStide) et ATP a permis de montrer l’activité de la kinase. L’analyse de deux inhibiteurs, staurosporine et compound C, ont permis de déterminer leur concentration d’inhibition IC50 comme étant 1 µM et 1.3 µM, respectivement. De façon surprenante aucune augmentation de l’activité de la kinase a été observée en incubant la kinase avec ou sans AMP ou ZMP, un analogue de AMP.

L'étude du rôle des sous-unités catalytiques TbAMPKα1/α2 in vivo ont révélé que le parasite est capable de survivre avec une absence partielle ou complète de ces deux sous-unités. En utilisant des souches knock-out (KO) de TbAMPKα1, nous avons

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observé que la croissance de la lignée n’était que partiellement influencée. Lorsque nous avons évalué l’ajout de l’effet de l’élimination de la deuxième sous-unité, TbAMPKα2, dans une souche knock-down nous avons observé une augmentation significative du temps de dédoublement, 9.75 ± 0.27 h, par rapport à la souche sauvage, 5.82 ± 0.09 h, sans observer de phénotype de mort cellulaire.

En conclusion, cette étude a permis d’identifier les sous-unités formant le complexe TbAMPK, de produire ce complexe in vitro et d’en débuter sa caractérisation constituant ainsi une première étape dans l’identification de potentielles nouvelles cibles thérapeutiques pour l’éradication du parasite.

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Summary of the thesis

Human African trypanosomiasis (HAT), also known as sleeping sickness, is a disease caused by the infection of Trypanosoma brucei (T. brucei), a unicellular parasite belonging to the order Kinetoplastida living in sub-Saharan Africa and transmitted by the tsetse fly of the genus Glossina. Two subspecies of T. brucei are responsible for the disease, T. b. gambiense responsible for the chronic form causing 97% of infections and T. b. rhodesiense responsible for the acute form causing the remaining 3% of infections.

Today, treatments for this disease are either very toxic, very complicated to administer or limited to the first phase (of infection) because of difficulties in crossing the blood- brain barrier needed to treat the second phase of infection. For these reasons, the discovery of new therapeutic targets and new drugs capable of eliminating T. brucei in its two stages of infection is very important to eradicate sleeping sickness. This thesis work was initiated after the discovery of a possible link between T. brucei adenosine kinase (TbAK) and the AMP-activated protein kinase cascade (AMPK). AMPK is a heterotrimeric complex known for its role in maintaining energy homeostasis in the cell by detecting the ratio AMP:ATP. It is composed of a catalytic subunit α which has to be phosphorylated in order to be activated and two regulatory subunits β and γ.

On the basis of this observation we decided to study T. brucei AMPK, a complex playing a primordial role in cell survival and very poorly characterized in T. brucei, particularly in the blood form of the parasite (blood stream form, BSF).

During this study, we confirmed the identity of the regulatory subunits TbAMPKβ (Tb927.8.2450) and TbAMPKγ (Tb927.10.3700), previously described in the procyclic form (characteristic form of the parasite in the vector) also in BSF parasites. In addition, we demonstrated that the two candidates of the catalytic subunit α (TbAMPKα1:

Tb927.3.4560, TbAMPKα2: Tb927.10.5310), identified by sequence alignment between the human α subunits and the parasite genome, do form a complex with TbAMPKβ and TbAMPKγ. This was possible by modifying the two-hybrid adenylate cyclase (BACTH) system to detect an interaction between two partners in a multiprotein complex. This new tool allowed us to demonstrate that the two candidates of the catalytic subunit form a complex with the other subunits.

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After identifying the subunits, the complex was expressed using a polycistronic expression system integrating the three subunits into the same plasmid under the control of a single promoter. This method has successfully produced the TbAMPKγβα2 complex in bacteria. The best yield was obtained after induction by 1 mM of IPTG of the protein production at 12°C for 72 h. Thanks to the presence of a His6-tag in N- terminus of the catalytic subunit, the complex could be isolated using Nickel affinity chromatography.

In order to improve the purity of the complex produced, an ion exchange chromatography and a size exclusion chromatography (SEC) were subsequently applied, allowing us to produce 0.16 mg of protein complex per g of initial bacteria mass. SEC-MALS (Multi angle light scattering) analysis confirmed that the absolute molecular weight of the purified complex was 160 kDa as expected.

Subsequently, the assessment of the phosphorylation state revealed that the complex expressed in the bacteria was already phosphorylated at the crucial phosphorylation site of the catalytic subunit, Thr172. In addition, an additional incubation with AMPK kinase did not increase the phosphorylation level. Similarly, incubation of the purified complex or production of the complex in the presence of a Lambda phosphatase (λPP) did not reduce the phosphorylation signal detected at the catalytic subunit.

The use of a bioluminescence assay to determine the level of ADP produced by the kinase reaction in the presence of ATP and a specific substrate (SAMStide) determined the EC50 of TbAMPKγβα2 to be 50.8 ± 6 ng/µL. Analysis of two inhibitors, staurosporine and compound C, determined their IC50 inhibition concentration to be 1 µM and 1.3 µM, respectively. Surprisingly no increase in kinase activity was observed by incubating kinase with or without AMP or ZMP, an AMP analogue.

The study of the role of catalytic subunits TbAMPKα1/α2 in vivo revealed that the parasite is able to survive with a partial or complete absence of these two subunits.

Using knock-out (KO) strains from TbAMPKα1, we observed that the growth of the line was only partially influenced. When we evaluated the addition of the effect of elimination of the second subunit, TbAMPKα2, in a knock-down strain we observed a significant increase in doubling time, 9.75 ± 0.27 h, compared to the wild strain, 5.82 ± 0.09 h, without observing a cell death phenotype.

In conclusion, this study allowed us to identify the subunits forming the TbAMPK complex, to produce this complex in vitro and to begin its characterization thus

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constituting a first step in the identification of potential new therapeutic targets for the eradication of the parasite.

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PART 1 Introduction

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1.1. Trypanosoma brucei

1.1.1. Trypanosoma classification

Trypanosomes are unicellular flagellate parasites belonging to the Kinetoplastida order of the Protozoan subkindom. Most trypanosomes are heteroxenous, which means that they need more than one host in their life-cycle, and need a vector to be transmitted.

The main diseases transmitted by parasite of this order are Leishmaniasis caused by different species of Leishmania, Chagas disease (also called American trypanosomiasis) caused by Trypanosoma cruzi and African trypanosomiasis (HAT;

also known as Sleeping sickness) caused by Trypanosoma brucei. The different mode of transmission between American trypanosomiasis and African trypanosomiasis is the characteristic that differentiates the two subgroups: stercoraria and salivarian.

Stercoraria species, such as T. cruzi develop in the intestinal tract of an invertebrate host and are transmitted via the secretion of its feces [1]. In contrast, African trypanosomes belong to the salivarian section of the genus Trypanosoma, which is characterized by transmission of the parasite via the saliva and the bite of the invertebrate host (tsetse fly, Glossina spp.) [1-3].

The salivarian group is divided into Duttonella (T. vivax), Nannomonas (T. congolense) and Trypanozoon (T. brucei, T. evansi and T. equiperdum) subgenera. Only parasites belonging to the Trypanozoon group, and more specifically the T. brucei species, are infective to humans. T. brucei is further divided into three subspecies: T. b. gambiense, T. b. rhodesiense and T. b. brucei, with the last one not being pathogenic for humans, but causing nagana disease in animals (African Animal Trypanosomiasis; AAT).

Trypanosomes are part of the “juxtaform” superclass which consists of trypomastigotes and epimastigotes and are defined by the presence of a flagellum that is laterally attached to the external cell body [1].

All three T. brucei species exhibit identical morphology and cannot be distinguished under the microscope. This fact causes serious problems with respect to anti-infective therapy since the treatment options are not the same for T. b. gambiense and T. b.

rhodesiense infections. Therefore, other techniques than microscopy or geographical observations are necessary to determine which parasite is the cause of the infection.

For this purpose, molecular marker for both infectious parasites were found, allowing

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a more precise differentiation. T. b. rhodesiense can be recognized by the presence of a serum resistance-associated (SRA) gene whereas T. b. gambiense by the presence of a TgsGP gene. The discovery of the TgsGP gene allowed the identification of a subgroup of T. b. gambiense called type 2 that does not harbor this marker. However, since type 2 T.b. gambiense was last isolated in 1992, it most likely does not contribute much to the spreading of the disease [4-7].

The resistance to infection of T. b. brucei is due to the presence of trypanosoma lytic factors (TLF1 and TLF2) in the human plasma [8, 9]. The TLF is composed of apolipoprotein A1, apolipoprotein L1 (APOL1) and a haptoglobin-related protein (HRP), and their antiparasitic activity is maximized when they are combined in one high density lipoprotein [10]. T. b. gambiense and T. b. rhodesiense have both acquired a specific mechanism to become resistant [11, 12]. T. b. rhodesiense avoids lysis by means of a Serum Resistance-Associated (SRA) protein, a truncated variant surface glycoprotein (VSG) present on the cell surface that binds to apolipoprotein L1 and neutralizes its action [4]. Humans that do not express apolipoprotein L1 have been shown to be sensitives to others infective species of Trypanosomes. T. b. gambiense, which does not contain SRA gene in its genome, escapes lysis in the human blood via with the help of a glycoprotein called TgsGP which is also a derivative of a VSG protein.

Knock-out experiment of TgsGP showed its importance in order to confer resistance to human serum to type 1 T. b. gambiense sub-species. More specifically it was shown to confer resistance against the trypanolytic protein APOL1 [13, 14]. In addition, L210S substitution in the haptoglobin-hemoglobin receptor (HpHbR) gene in T. b. gambiense resulted in a reduction in the binding to APOL1 and thus a reduction in its internalization conferring TLF1 resistance to the parasite [13, 15].

1.1.2. Morphology and cell structure

African trypanosomes are unicellular parasites that live in body fluids such as blood, lymph or cerebrospinal fluid. Their size is about 20 µm long and 5 µm wide, and their general structure and organelles are shown Figure 1. Trypanosomes have two DNA- containing organelles, the nucleus and the kinetoplast, the latter being the characteristic organelle of the Kinetoplastida order. Glycosomes (another organelle specific to the kinetoplast organisms) are considered to have evolved from peroxisomes and contain the first seven steps of the glycolytic pathway among other

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biochemical pathways [2]. The parasites harbour a single unit of several eukaryotic organelles, such as the Golgi apparatus and the mitochondrion.

Trypanosomes are characterized by the presence of a single flagellum composed of the conventional 9+2 axonemes. The flagellum is attached at the basal body and exits the cellular membrane at the level of the flagellar pocket (which is important also for the interaction with the environment). The flagellum plays a role in the mobility of the cell as well as in mechanically sweeping surface material. Flagellar movement can induce endocytosis of antibodies bound to the cell surface [2, 16].

Figure 1 General structure of an African trypanosome (Image taken from [2]) and Trypanosomes among blood cells (Image taken from [12]).

A very dense coat of glycoproteins (the so-called Variant Surface Glycoproteins;

VSGs) covers the surface of trypanosomes (thickness: 15nm). These glycoproteins (or VSGs) are the predominant antigen present at the surface of the parasite. VSGs are very immunogenic and the immune response is usually focused on these antigens.

However, by the time the immune response is ready to attack the trypanosomes carrying the specific glycoproteins, some of them will already have changed the type of VSGs at its surface, thus escaping the response of the immune system [12, 17, 18].

This is possible because the T. brucei genome contains up to 2000 different genes and pseudogenes coding for VSGs, but only one type is expressed at a time [12, 19].

In addition, it has been observed that even more variants of VSG genes are obtained by exchange of small region between two genes, thus creating new chimeric variants.

The VSG coat is specific to the bloodstream form of the parasite, and its expression needs to be activated already in the salivary gland of the tsetse fly before injection into

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the human host [20]. In the insect, another type of proteins, called procyclins, composes the coat.

Finally, the presence of transport proteins, called permeases, in the plasma membrane allows the parasite to access all the nutrients from the host needed for surviving [21, 22].

1.1.3. Vector

T. brucei are transmitted by the bite of a tsetse fly belonging to the genus Glossina which is part of the Glossinidae family of the Diptera order. Both female and male of this genus are haematophagous and possible vectors of African trypanosomiasis. The Glossina genus has been divided into 3 subgenera (Nemorhina, Glossina s. str. and Austenina) all together divided into 31 species and subspecies. Their classification is based on external characteristics, the shape of genitals and the geographical distribution. Depending on the infected tsetse fly species, it shows different abilities of transmitting T. b. gambiense or T. b. rhodesiense diseases [12]. Their distribution, with an area of 10 million km², is concentrated in Sub-Saharan African countries [2, 23].

Figure 2 shows the picture of two main tsetse fly subspecies, Glossina palpalis (G.

palpalis) and Glossina morsitans morsitans (G. morsitans morsitans).

Figure 2 Tsetse fly. (A) Female Riverine (G. palpalis). (B) Female Savannah Tsetse fly (G. morsitans morsitans).

(C) Bloodfed female Savannah Tsetse fly (G. morsitans morsitans). Pictures taken from[24]

Subgenus Nemorhina (“riverine Tsetse”)

Species of this subgenus are of small (6-8 mm) to medium (8-10 mm) size and are found in West and Central Africa in vegetation located close to water. Moreover, evidence of peri-urban transmission in medium and large towns has been observed largely extending the tsetse fly distribution. The subspecies G. palpalis and G. fuscipes are considered to be the main vectors of both HAT and are more and more found to

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be responsible for AAT due to demographic growth and their ability to adapt to high density population regions [2, 25].

Subgenus Glossina s. str. (“savannah tsetse”)

Species of this subgenus are of medium size (8-11 mm) and are found principally in savannah woodland. They are the main vector of AAT as they are found in region with a high density of wild animals. Some species of this subgenus, principally G. morsitans, G. pallidipes and G. swynnertoni, are also considered as important vectors for rhodesiense HAT [2].

Subgenus Austenina

Species of this subgenus are of large size (11-16 mm) and are mainly found in forest belts. They could be good vector for AAT but only few animals live in the region where these tsetse flies are found [2].

Due to environmental changes throughout the year (rainy/dry and warm/cold seasons) the density of tsetse flies varies extensively. Tsetse flies can only survive in tropical areas because they need between 50-60% of humidity for the savannah subspecies and between 65-85% for the riverine and forest species. Moreover, the temperature necessary for their survival is between 16 and 36 °C and higher temperature can be lethal for both pupae and adult flies. During the short life of the tsetse fly the female produces one at the time about 3 to 5 larvae only. For this reason, the growth of a tsetse population is very limited and small interferences can easily affect the population density. For example, the risk of being killed during the blood meal could represent a real threat to the population. To avoid being killed, the tsetse fly tends to bite where it is protected from possible defensive movements of the host [2].

During a blood meal the tsetse fly first injects some saliva into the skin which provokes a vasodilatory effect and also prevents blood clotting. It is at this stage where an infected fly transmits matures trypanosomes to the new host. Since an infected tsetse fly will remain a carrier of trypanosomes its whole lifetime, control and eradication of the vector is very important to reduce the risk of HAT and AAT transmission.

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1.1.4. Life-cycle

The life-cycle of trypanosoma, which is complex, is divided into two major stages, one in the blood of a mammalian host and the other one in an insect host, the tsetse fly.

They are called blood stream form (BSF) and procyclic form (PCF), respectively.

Trypanosomes, extracellular parasites, need to undergo a series of transformations, such as modification of cell shape, metabolism and gene expression, during their life- cycle in order to adapt to two very different environments (Figure 3) [26]. The cycle starts with a blood meal of an infected tsetse fly which will result in injection of metacyclic trypomastigotes, the infective form, in the blood of the mammalian host.

Metacyclic trypanosomes are adapted to survive in mammalian blood because of the presence of a thick surface coat composed of glycoproteins. Once injected metacyclic trypanosomes will proliferate at the site of infection. After entering the lymph and the blood they differentiate into proliferating long slender bloodstream form which will multiply by binary fission. In the second stage of the infection trypanosomes will cross the blood brain barrier and invade the central nervous system (CNS) [2].

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Figure 3 Life cycle of Trypanosoma brucei. (1) Ttsetse fly takes a blood meal and injects metacyclic trypomastigotes.

(2) Injected metacyclic trypomastigotes transform into bloodstream trypomastigotes, which are carried to other sites.

(3) Trypomastigotes multiply by binary fission in various body fluids (e.g. blood lymph and spinal fluid). (4) trypomastigotes in blood. (5) Tsetse fly takes a blood meal, bloodstream trypomastigotes are ingested. (6) Bloodstream trypomastigotes transform into procyclic trypomastigotes in tsetse fly’s midgut. Procyclic trypomastigotes multiply by binary fission. (7) Procyclic trypomastigotes leave the midgut and transform into epimastigotes. (8) Epimastigotes multiply in salivary gland. They transform into metacyclic trypomastigotes. Image taken from [27]

In order to complete the cell-cycle and to be taken up by the tsetse fly, trypanosomes need to differentiate into the non-proliferating stumpy form in the bloodstream of the mammalian host. As shown in Figure 4 the long slender form proliferates and increases the parasitaemia in the mammalian host. This increase induce accumulation of stumpy induction factor (SIF) a chemically uncharacterized signal, that will induces the transition from the long slender to the stumpy form at the peak of parasitaemia [28].

The stumpy form, which still expresses VSG on its surface, is preadapted to survive in the tsetse fly and can be collected by the insect during a blood meal. In insects it transforms into the procyclic form that expresses procyclins on the surface (EP/GEET)

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instead of VSG. Since stumpy form parasites do not proliferate anymore, they do not change the VSG coat, thus become accessible and are killed by the immune response of the mammalian host. Only few revert to long slender and proliferating parasites that are capable to escape the immune response by switching the VSG gene expressed at the surface, and will restart the cycle [29, 30]. This fluctuation in parasitaemia is considered necessary to avoid fast and lethal damage to the host [30, 31].

Figure 4 Life-cycle stages of Trypanosoma brucei Bloodstream form. During the course of parasitaemia the long slender form proliferates and increases the infection. As the parasitaemia increases the accumulation of a stumpy induction factor (SIF) induces the transition from long slender proliferating form to stumpy non-proliferating form that is pre-adapted at the life in the insect. Both forms still express VSG at their cell surface. In the tsetse fly the parasite transforms into procyclic form expressing procyclins (EP/GPEET) instead of VSGs at their cell surface. If not taken up by the tsetse fly the majority of the stumpy form parasite is either killed by the immune response of the host or will degenerate in the following days lowering the parasitaemia. Only few parasites will revert to the proliferative long slender form escaping the immune response by changing the VSG gene expressed at their cell surface. Figure taken from [29].

Once in the midgut of the fly, the stumpy form parasite differentiates into procyclic trypomastigotes. The shock of temperature, from 37°C to 20°C, induces the switch from VSG to procyclins expression at the surface of the stumpy form parasite. Before maturation to the infective metacyclic trypomastigote form the parasite will move to the salivary gland and transform into the epimastigotes form. The maturation step can take between 18 and 35 days [2, 20, 32].

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In the tsetse fly it has been shown that sexual reproduction is possible but not mandatory. This process will result in exchange of genetic information that will allow rapid adaptation and evolution for example by gaining resistance to a new drug.

The tsetse fly needs to take a blood meal from an infected mammal in order to be a carrier for the disease, no newly-born flies have been reported to be already carrier of the disease. Only about 0.1% of infected flies are carrier of the mature infective form of the parasite, this is due to the fact that the maturation cycle of the parasite is only rarely completed in nature, but once the fly is infected it will remain so for the rest of its lifespan (on average 3 weeks for the male and 35 days for the female tsetse fly) [12].

1.1.5. T. brucei Genome

The parasite genome, of approximately 35 Mb per haploid genome, was completely sequenced in 2005 and its sequence can be found on GeneDB (http://www.genedb.org/Homepage/Tbruceibrucei927) website and TritrypDB (http://tritrypdb.org/tritrypdb/) website [2, 33].

As mentioned before, the parasite contains two major DNA carrying organelles: the nucleus and the kinetoplast. The nucleus is composed of 11 diploid maxichromosomes (1-6 Mb), several intermediates chromosomes (300-900 kb), and many minichromosomes (50-100 kb) that contain the information for the numerous VSG genes.

The kinetoplast contains the mitochondrial genome which is particularly large. It is composed of several circular DNA structures that can be separated in maxi-circles (20kb) coding for proteins, and mini-circles (1-2 kb) coding for guide RNAs that are needed for RNA editing which usually consists in addition and more rarely deletion of U-residues to the newly transcribed RNAs [34]. Studies on the kinetoplast genome have shown that many of the genes after being transcribed into RNAs, need a very complex editing process which is specific to Kinetoplastid protozoa in order to form functional mRNA [2, 35].

Gene transcription of trypanosomatids differs from other eukaryotes. Several genes are constitutively transcribed as a single segment of RNA (polycistronic construct) that needs further processing in order to be stabilized. Trans-splicing process, addition of

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a poly-A tail at the 3’ and a spliced leader and 5’ terminus are necessary to form mature mRNA (Figure 5) [2, 36].

Figure 5 Example of transcription and post transcription regulation in trypanosomatids. An example of polycistronic transcription representing three gene clusters (PGC1-3). Transcription of cluster 2 include 3 genes that are transcribed together and processed by polyadenylation and trans-splicing in order to form mature mRNA. The trans- splicing process add a capped SL RNA (shown as a yellow square in the image) in 5’ of each mRNA, whereas the poly-A-tail is added in 3’ of the mRNA. Figure adapted from [37].

The capability of the parasite to adapt to different environments during its life-cycle is possible by significant changes in gene expression that occur at the post- transcriptional level by regulating mRNA and protein levels. In addition to post- transcriptional modifications and polyadenylation, RNA export and stability, protein translation and stability are also very important regulators [38-40].

This shows the importance of the translation regulation step in order for the trypanosome to adapt to different environments such as mammals and insects [2].

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1.2. Human African trypanosomiasis (HAT)

1.2.1. The disease

Human African trypanosomiasis (HAT), also known as sleeping sickness, is a disease present in sub-Saharan Africa [12] and is caused by two of the three subspecies of T.

brucei. T. b. gambiense is responsible for the gambiense HAT, the chronic form of the disease, whereas T. b. rhodesiense is responsible for the rhodesiense HAT, the acute form of the disease. Even though both parasite infections are named HAT diseases, in reality the diseases transmitted are very different at the biological, clinical, therapeutical, geographical and epidemiological levels [41]. Symptoms are similar for both diseases but a difference can be observed in their frequency and severity.

The third subspecies, T. b. brucei, is the causative agent of nagana in domestic and wild animals (Animal African Trypanosomiasis, AAT) but it is not pathogenic for humans. [12] Nagana was associated with Trypanosoma infection in 1899 by David Bruce and further studies confirmed the infection of domestic animals was the result of a bite of a tsetse fly that had been infected while blood-sucking infected wild animals.

[16, 42] T. b. brucei will cause fever, weakness and lethargy in animals, leading to weight loss and anaemia and usually to death if untreated.

The infection is transmitted mainly by the bite of an infected tsetse fly, however, other ways of transmission have been observed. It has been shown that trypanosomes can cross the placenta and infect the foetus, they can be transmitted from other insects or during sexual contact, and finally accidental transmission in the laboratory with contaminated needles were reported. [43]

Figure 6 and Figure 7 shows the geographical distribution of the two forms of HAT.

Gambiense HAT, which is more present in central and West Africa, is considered endemic in 24 countries and it is responsible for 97% of the disease. Rhodesiense HAT, more spread in East and southern Africa, is considered endemic in 13 countries and it is responsible for the remaining 3% of the disease [12, 43].

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Figure 6 Distribution and incidence of human African trypanosomiasis. The black line divides the regions between the predominance of T.b. gambiense and T.b. rhodesiense. Image taken from [12]

Despite the distribution of HAT is usually well separated between gambiense and rhodesiense, it has been observed that in Uganda both form of HAT are present. Since the treatment is different depending on the parasite, this observation raises the problem of determining co-infection and implementing co-treatment [42, 44, 45].

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Figure 7 Maps of distribution of Human African trypanosomiasis reported in 2016 for T. b. gambiense (A) and T.b.

rhodesiense (B). Figure adapted from [66].

The disease progression can be divided into two principal stages, the first called haemo-lymphatic stage and the second called meningo-encephalitic stage.

The haemo-lymphatic stage, as the name suggests, is the stage where the parasite is found in the blood, in lymph nodes, in several systemic organs (such as spleen, heart and liver) and other organs (such as eyes and endocrine related organs) [46]. The second stage of the disease starts once the parasite crosses the blood-brain barrier (BBB) and enters the CNS. It is during this second stage that all neurological symptoms occur, such as sleep disturbance (that gave the name to the disease). However, several signs occur during both stages and are more or less frequent within individuals and geographical foci of infections.

The infection by either parasite leads to death if not treated, but the duration of T.b.

gambiense infection, the chronic progressive form of the disease, is estimated to be

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around 3 years [47] whereas the duration of T.b. rhodesiense infection, the acute form, is between few weeks to 6 months maximum [48].

Gambiense HAT: First stage of gambiense HAT can easily be unnoticed since the symptoms can be intermittent, not severe and will only increase during the second stage. Some of the symptoms are lymphadenopathy (usually growth of the posterior cervical lymph node), intermittent fever, progressive headache, pruritus and electrocardiographic alterations (increase in second stage of infection). Once the parasite crosses the BBB and the CNS is attacked the principal symptom observed is the sleeping disorder. The parasite induces a dysregulation of the circadian rhythm and the patient will experience fragmentation of the sleeping pattern. Some psychiatric symptoms are already present in the first stage of the disease, but will increase in intensity during the second stage, and new symptoms depending on the injured region of the brain can occur. One common symptom is the deep sensory disturbance that induces strong pain even for a small shock. In the terminal stage of the disease severe dementia, epilepsy and severe disturbance of consciousness are observed and will lead to the death of the patient [2, 49, 50].

Rhodesiense HAT: Symptoms of rhodesiense HAT are similar to gambiense HAT, however some differences can be observed. Compared to gambiense HAT, it has been noticed that the presence of chancres at the site of infection are more frequent in rhodesiense HAT. The lymph node that are enlarged are more the submandibular, the axillary and the inguinal instead of the cervical one. Moreover, for unknown reasons, differences in symptoms severity and emergence are observed between foci [2, 51, 52].

1.2.2. Epidemiology and current situation

Human African Trypanosomiasis (HAT) is a neglected disease localized in sub- Saharan Africa, impacting 70 million people. Its geographical distribution is extremely focal due to the necessity for the tsetse fly to find the good environment [2, 53].

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In the last century, several epidemics and remissions were observed. When it was discovered that trypanosome was the cause of HAT, the parasite had already been responsible for the deaths of hundreds of thousands of people between 1896-1906 [12, 16, 44] and again in 1920. The social and economic development that led to the mobility of the population and animals also contributed to the spread of the disease [2].

In the 1960s, thanks to various control measures, HAT was finally considered under control and only a few new cases had been reported. Important mass killings of animals (responsible for being a reservoir of the vector), eradication of wooded regions in sub- Saharan Africa, better control of insect vector, systematic screening to improve control of possible infected populations and finally better access to effective chemotherapy were the important steps followed to achieve this result. However, in 1998, due to various factors, such as war, famine and the resulting instability, which led to a reduction in the control of possible infected areas, according to the WHO the number of reported cases increased again to 300,000 [12, 16, 54, 55].

Following this big resurgence of the disease, efforts by the WHO and other partners resulted in a drastic reduction of reported cases in 2006 (around 50’000-70'000) and less than 10'000 by 2009 [54, 56]. In 2011 WHO reported 6'631 new cases [57].

However, this result may not completely correspond to the reality as many cases may have been missed, for example due to difficulties of reaching very remote areas and absence of equipped facilities to recognize and treat HAT [44]. The collaboration between several non-governmental organizations (NGOs), WHO and two private pharmaceutical companies, Sanofi-Aventis (for pentamidine, melarsoprol and eflornithine) and Bayer HealthCare (for Suramin and nifurtimox), could reduce to

<3’000 of gambiense HAT cases in 2015 thanks to the free distribution of treatment in endemic areas [58]. This partnership created in 2001 was renewed in 2006, 2011 and again in 2016. In 2016 the WHO observed that the newly reported cases of T. b.

gambiense and T. b. rhodesiense dropped from 27’862 and 619 in 1999 to 2131 and 53 in 2016 for the two forms of the disease, respectively. The number of actual cases is estimated to be around 20’000 with 65 millions of people at risk and 13 millions living in areas at moderate/high risk of infection [43].

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Domestic and wild animals play an important role as carriers and reservoirs for both pathogenic parasites. However, T.b. gambiense is an anthroponotic disease, meaning that it depends mostly on human-to-human transmission where men are considered the main reservoirs [59]. The risk of being infected depends largely on the activity exerted during the day and the environmental characteristics. For example, when for agricultural expansion reasons the tse-tse territory is reduced an increase of new infections due to the approach of man to the infected regions is observed [2].

T.b. rhodesiense is a zoonotic disease, which means that the disease is usually transmitted from a vertebrate, mainly cattle, to a human [60, 61]. It has been shown that for rhodesiense HAT several animals play a role as reservoirs or hosts, however observations have not been made on a large scale, on wild animals, due to difficulty of collecting blood samples [2]. The risk of infection is higher for adults than for children when their daily activity includes working outside; like, farming, collecting firewood, fishing, hunting or washing clothes. The risk is even increased when wildlife is close to the human habitats or livestock since they can act as reservoirs [2, 62].

A very important tool used to study and predict HAT evolution was the creation, in collaboration with WHO Member States, of an Atlas of human African trypanosomiasis [63-65]. The Atlas, which maps the distribution of the reported cases at the village level, can be used as a start for analysing the evolution and changes in HAT infections and distribution. The map is continuously updated with observations made in the field [2, 43]. Factors that will change the geographic distribution of HAT are population growth and climate changes. In fact, the population growth can result in destruction of tsetse fly habitats which results in new distribution of their foci of action. Improvement in the collection of data from population growth and geographical distribution will, thus, be very important [2].

Murine studies showed the importance of the host genotype as well as the impact of co-infection in the severity of the disease outcome [67, 68]. It has been shown that coinfection of both acute and chronic form of the disease resulted in a reduction of the infection in the early stage probably due to the competition for common resources, allelopathic interference and immune-mediated competition. Moreover, Balmer et al.

observed that coinfection with a less virulent strain reduces the density of the more infectious parasite and therefore increases the surviving rate of the host [68]. These

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observations are very important, especially considering the problem Uganda faces due to the overlap of the two infections.

1.2.3. Diagnosis and treatment 1.2.3.1. Diagnosis

Parasitaemia is very different between T.b. gambiense and T.b. rhodesiense infection.

In the first case it is usually very low (<100 parasite per mL) which is very difficult to observe under microscope, whereas in the second case the amount of trypanosome is usually higher. Moreover parasitaemia usually fluctuates during the infection due to the immune response of the infected host and the ability of the parasite to evade the immune response by quickly exchanging the antigen present on the surface, rendering the detection more complicated Figure 4 [2].

The diagnosis and the treatment of the diseases caused by the two parasites form are very different. Observation of early symptoms such as swollen lymph node, repeated fever and neurological signs are not enough to start a treatment but can induce the suspicion in order to further investigate the patient. Moreover, since serological tests are also not 100% reliable, it is still important to confirm the infection by observing parasites in the body fluid.

Several serological tests are available to detect gambiense HAT. However, depending on feasibility, time consuming, material requirements and costs, they can be applied directly in the field (usually rural regions of Africa), or only in specific laboratories with specific equipment.

Most of these tests rely on the presence of two antibodies directed against two specific antigens (VSG antigen type LiTat 1.3 and 1.5) of T. b. gambiense. The card agglutination test (CATT) is a very fast, inexpensive and simple test developed in the late 1970s with a sensitivity around 87-98% and a specificity around 93-95%. It has been largely used in all control programs for serological screening of the population [69-71]. However, its use in remote areas is difficult due to the need of access to electricity and to continuously maintain the kit at cold temperatures. Several other tests relying on the same detection of specific antigens, using ELISA based immunofluorescence assays, also need expensive equipment, like fluorescent

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microscopes and require electricity, high amounts of water or specific storing conditions that cannot be ensured in the field [2]. A recent test called Rapid Diagnostic test (RDT) overcomes all these problems. RDT is an immunochromatographic test that detects specific antibodies of a T. b. gambiense infection from a fresh blood drop in only 15 minutes without the need of specific equipment or specific storage conditions.

In September 2017 the second generation of RDT test (SD BIOLINE HAT 2.0) developed by FIND and supported by the Bill & Melinda Gates Foundation, UK aid from the UK government and the Swiss government was commercially launched [72].

In order to confirm the presence of the parasite, the blood or lymph fluid of a patient being positive at the serological or clinical level need to be observed under the microscope. However, these observations are very time-consuming and due to the usually low parasitemia of gambiense HAT, false-negative results occur very often. For this reason, absence of observation of trypanosoma under a microscope cannot completely exclude the possibility of an infection [2].

In some cases, molecular detection of trypanosoma DNA or RNA can be performed in order to confirm the presence of the parasite. These tests can be performed also on stored samples and are therefore a very interesting test to perform when live parasite detection fail to demonstrate the presence of parasites. Molecular detections rely on the specificity of one single copy gene, the TgsGP, which if detected on agarose gel after PCR amplification, will confirm the presence of T.b. gambiense [4, 73, 74].

In the case of rhodesiense HAT due to high antigenic variation (higher compared to the gambiense subspecies) CATT screening cannot be used as there is no specific VSG gene that can be used for diagnosis. For molecular detection the technique is based on the detection of the SRA gene which is specific to this subspecies. However, compared to gambiense HAT, rhodesiense infection usually results in a much higher parasitaemia (up to 10 000 trypanosomes/ml), so microscopic detection of parasite is much easier [2].

In order to be able to give to the patient the right treatment, it is mandatory to know the species and also of the stage of infection. In both cases, analysis of the cerebrospinal fluid (CSF) for detection of trypanosomes is necessary [75]. Since lumbar puncture is a very invasive technique it is only performed once parasitemia has already been

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demonstrated or when strong clinical or serological sign are present. Before performing lumbar puncture, in particular if a T. b. rhodesiense infection is suspected, a dose of suramin is administered upfront in order to clear the blood and reduce the risk of introducing the parasite in the CSF during the manipulation. The CSF is then analysed for the presence of white blood cells and/or parasites in order to establish the progress of the disease [2].

Diagnosis outlook: Simple and rapid tests available to use in the field have been a huge progression in the control and screening of HAT. Improvement in existing test in order to render them more sensitives, less expensive and easy to use in the field are some of the concern that are taken into account for the future [76]. One important progress has been the development of Loop-mediated isothermal amplification test (LAMP) [77]. This technique relies on the detection and amplification of a specific DNA sequence by a set of specific primers used at an amplification temperature between 60-65°C. The result can be observed by simple observation in the colour change in the mixture, and the specificity and sensitivity has been proven to be very high. The adaptation to detect the specific gene marker TgsGP and SRA for gambiense and rhodosiense HAT, respectively, is in progress. Cost-efficacy compared to standard methods and sensitivity level in patient samples need to be proven before the implementation of the test in the field [2, 74].

Since 2009 a bank collecting blood, serum, CSF, saliva and urine from infected and uninfected patient from endemic areas was created by the WHO and made accessible to researchers in order to improve the diagnostic tools. [43]

1.2.3.2. Treatment

The early detection of an infection is very important to avoid disease progression to the second stage for which the therapy is risky and often unsuccessful [43]. At the present time only 5 drugs are available for fighting HAT. The currently used drugs were mostly discovered in the first half of the 20^th century. All of these drugs have serious disadvantages as they are very toxic, difficult to administer or may not cross the BBB which makes them only active against the treatment of the first stage of the disease.

For this reason, it is crucial to know the stage and the parasite subspecies to choose the right drug treatment. [2, 12].

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The drugs currently used and chemotherapeutic treatment of HAT are eflornithine, melarsoprol, nifurtimox, pentamidine and suramin (Figure 8 and in Table 1)

Figure 8 Chemical structures, first year of publication of drugs use for HAT treatment and brand name.

Pentamidine: This molecule was discovered in 1940 and is used for the treatment of the first stage of T.b. gambiense infection. The mode of action of the pentamidine is unknown, however, some possible mechanisms such as binding to nucleic acids, disruption of kinetoplast DNA, inhibition of RNA-editing and inhibition of mRNA transplicing or inhibition of the plasma membrane CA²⁺-ATPase have been hypothesized [78, 79]. The treatment with pentamidine consists in one intramuscular injection per day over one week. The reported adverse reactions to the treatment are site pain, transient swelling, abdominal pain, gastrointestinal problems and hypoglycaemia (in less than 40% patients) [80-82].

Suramin: Suramin was discovered in 1920 and is used to treat first stage of T.b.

rhodesiense infection. However, the treatment is long, needing 5 slow intravenous injections (after the first test injection), one every 7 days. Nephrotoxicity, peripheral neuropathy and bone marrow toxicity accompanied by agranulocytosis and thrombocytopenia are possible rare and reversible adverse reactions to the drug [12].

As for pentamidine, the mode of action of suramin is unknown.

Melarsoprol: Melarsoprol was discovered in 1949 and is the only drug being active against second stage of T.b. rhodesiense infection, but can also be used for second stage of T.b. gambiense infection, when a different treatment regimen is considered.

Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of Trypanosoma brucei

Thesis

Reference

Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of Trypanosoma brucei

Identification and analysis of the putative AMP-activated protein kinase (AMPK) in blood stream form of

Trypanosoma brucei

Acknowledgments

Table of contents

Abbreviations

Résumé de la thèse

Summary of the thesis

PART 1

Introduction