Ghent University Faculty of Veterinary Medicine

Home , Trypanosoma congolense, Trypanosoma suis

GHENT UNIVERSITY

FACULTY OF VETERINARY MEDICINE

Academic year 2015-2016

THE IMPORTANCE OF WILDLIFE AS A RESERVOIR FOR HUMAN AND ANIMAL AFRICAN TRYPANOSOMIASIS

Kim VAN DE WIEL

Promoter: Prof. Dr. Pierre Dorny Literature Review Co-promoter: Prof. Dr. Louis Maes as part of the Master’s Dissertation

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GHENT UNIVERSITY

FACULTY OF VETERINARY MEDICINE

Academic year 2015-2016

THE IMPORTANCE OF WILDLIFE AS A RESERVOIR FOR HUMAN AND ANIMAL AFRICAN TRYPANOSOMIASIS

Kim VAN DE WIEL

Promoter: Prof. Dr. Pierre Dorny Literature Review Co-promoter: Prof. Dr. Louis Maes as part of the Master’s Dissertation

PREFACE

For my dissertation, I had the pleasure to choose my own subject. As an enthusiast of parasitology with a keen interest in zoonotic diseases, and a love for Africa, I can’t imagine any other topic that would have combined these aspects as well as this one.

First of all, I would like to thank my promoter, Prof. Dr. Pierre Dorny, who immediately replied with a positive message when I asked if I could write about ‘the animal reservoir of sleeping sickness’. I would also like to thank my co-promoter Prof. Dr. Louis Maes, whose course material had made me enthusiastic about tropical parasites in the first place. Thanks to both of them I had the freedom to fill in this dissertation to my own liking. Also a big thanks for suggesting articles, helping me find them, and reviewing my dissertation in time, even though it was very last minute from my side.

In the second place I would like to thank my parents for their patience and unconditional support during the first, but definitely also the final years of veterinary school.

Last, I would like to thank my friends. Some of them for their advice on how to start writing, others for reviewing some of my work. But most of all, I would like to thank the friends who told me to stop complaining and continue on, whenever I had tiny emotional breakdowns about unimportant things.

TABLE OF CONTENT PREFACE

TABLE OF CONTENT

SUMMARY ...... 1 SAMENVATTING ...... 2 INTRODUCTION ...... 3 LITERATURE STUDY ...... 4 1. The parasite: Trypanosoma ...... 4 1.1. Morphology ...... 4 1.2. Life cycle...... 4 1.3. Classification ...... 6 2. The vector: Glossina ...... 7 2.1. General features ...... 7 2.2. Distribution ...... 7 2.3. Taxonomy and subgenera ...... 7 2.4. Feeding preferences ...... 8 2.5. Important vector species ...... 10 3. Human African trypanosomiasis ...... 12 3.1. Causative agents ...... 12 3.2. Clinical symptoms ...... 12 4. Animal African trypanosomiasis...... 14 3.1. Causative agents ...... 14 4.2. Clinical symptoms ...... 14 5. Reservoirs ...... 16 5.1. Reservoir for human African trypanosomiasis ...... 16 5.2. Reservoir for animal African trypanosomiasis ...... 20 5.3. Control of reservoirs ...... 21 DISCUSSION ...... 23 REFERENCES ...... 24 ANNEXES ...... 30

SUMMARY African trypanosomiasis is an infectious disease that affects both people and animals. It is caused by small protozoa that can be transmitted to humans and animals via hematophagous insects. Several species of trypanosomes have been identified as pathogenic in vertebrate hosts. The two subspecies, Trypanosoma brucei gambiense and Trypanosoma brucei rhodesiense, are responsible for causing sleeping sickness in humans. In West Africa, T. b. gambiense seems to be the causative agent of the chronic form of the disease, whereas T. b. rhodesiense ensures a more acute onset of the disease in East Africa. Multiple other trypanosomes are held accountable for the disease complexes caused in domestic animals. The most economically important trypanosomes in livestock are T. congolense and T. vivax, which cause nagana in cattle.

These pathogenic trypanosomes are restricted to the African continent by their vector, the tsetse fly. As the only cyclical vector of the trypanosomes, the abundance of these flies in sub-Saharan Africa results in a large amount of countries at risk of trypanosome infection. Other hematophagous insects are capable of transmitting these parasites mechanically. For some trypanosome species, like T. vivax, this assures their spread beyond the tsetse belt.

Although control measurements for human sleeping sickness take place on a large scale, persistence of the disease in several regions has been observed. A possible animal reservoir for T. b. gambiense and T. b. rhodesiense has been suggested very early on. The feeding preferences of the tsetse fly indicate that these insects feed on, and possibly also infect, a great variety of hosts. It was revealed that both T. b. gambiense and T. b. rhodesiense had a reservoir in several domestic and wildlife species, but the importance of these reservoirs in the epidemiology of the disease is still unclear.

T. congolense and T. vivax continue to have a big impact on livestock in Africa. These trypanosomes appear to be more widespread than their human-infective cousins, as case detecting in livestock is not executed on such a large scale as is done for sleeping sickness. A wildlife reservoir seems to be the reason for the spread of these trypanosomes, but the role of wild animals in the maintenance of animal trypanosomiasis is even less understood than in human sleeping sickness.

Keywords: Trypanosoma – Nagana – Sleeping sickness – Reservoir – Wildlife

SAMENVATTING Afrikaanse trypanosomiase is een infectieuze ziekte die zowel mensen als dieren treft. De ziekte wordt veroorzaakt door protozoa, die door bloedzuigende insecten naar mens en dier kunnen worden overgedragen. Bepaalde trypanosomen worden als pathogeen beschouwd in hun gewervelde gastheren. De twee ondersoorten, Trypanosoma brucei gambiense en Trypanosoma brucei rhodesiense, zijn verantwoordelijk voor het ontstaan van slaapziekte bij de mens. In West-Afrika veroorzaakt T. b. gambiense de chronische vorm van trypanosomiase, terwijl T. b. rhodesiense in Oost Afrika aanleiding geeft tot een meer acute vorm van de ziekte. Meerdere trypanosomen zijn verantwoordelijk voor de ziektecomplexen die worden teruggevonden in gedomesticeerde dieren. T. congolense en T. vivax, die nagana veroorzaken in runderen, zijn de economisch meest belangrijke soorten voor de veestapel.

Deze pathogene trypanosomen worden door hun vector, de tsetse vlieg, geografisch gelimiteerd in hun verspreiding en zijn enkel te vinden op het continent Afrika. De tsetse vlieg is de enige cyclische vector van deze parasieten, en door de overvloedige aanwezigheid van de vliegen in sub-Sahara Afrika, is het risico op infectie in veel Afrikaanse landen aanwezig. Trypanosomen kunnen via andere bloedzuigende insecten mechanisch verspreid worden. Dit is de reden dat sommige soorten, zoals T. vivax, tot ver buiten de tsetse gebieden gezien worden.

Hoewel controlemaatregelen tegen slaapziekte bij de mens op grote schaal worden ondernomen, blijkt de ziekte in verschillende gebieden te persisteren. Een mogelijke dierlijk reservoir voor T. b. gambiense en T. b. rhodesiense werd al vrij vroeg gesuggereerd. Onderzoek naar de voedselvoorkeuren van de tsetse vlieg heeft aangetoond dat deze insecten diverse gastheren bijten voor hun bloedmaal. Dit kan mogelijks resulteren in de verspreiding van trypanosomen onder een grote groep verschillende gastheren. Zowel T. b. gambiense als T. b. rhodesiense bleken een reservoir te hebben binnen verschillende gedomesticeerde en wilde dieren. De rol van deze reservoirs in de verspreiding van de ziekte is echter nog niet helemaal opgeklaard.

T. congolense en T. vivax hebben een grote impact op de veestapel in Afrika. Deze trypanosomen blijken meer verspreid te zijn dan de soorten die mensen infecteren, doordat controlemaatregelen voor dierlijke trypanosomen minder fel zijn uitgebouwd dan de controlemaatregelen voor slaapziekte. Een reservoir in wilde dieren bleek een reden voor de verspreiding van deze trypanosomen, maar de rol van wildlife in de epidemiologie van dierlijke trypanosomiasis is zelfs nog minder begrepen dan hun rol in de verspreiding van slaapziekte.

Sleutelwoorden: Trypanosoma – Nagana – Slaapziekte – Reservoir – Wildlife

INTRODUCTION Trypanosomiasis is a disease caused by the parasitic protozoan Trypanosoma. Although these parasites can cause disease in many different hosts, the pathogenic, tsetse-transmitted trypanosomes of Africa are responsible for transmitting human African trypanosomiasis (HAT) or sleeping sickness, and animal African trypanosomiasis (AAT).

Sleeping sickness in humans is caused by two species of the subgenus Trypanozoon: T. b. gambiense and T. b. rhodesiense. According to the WHO (2013), around 70 million people are annually at risk of getting infected with these human-infective trypanosomes. Trypanosomiasis does not only cause severe suffering and mortality in humans, but it also affects their livestock significantly. T. congolense and T. vivax are among the many trypanosomes that can infect animals. Approximately 60 million cattle in sub- Saharan Africa are at risk of contracting trypanosomes (Kristjanson et al., 1999). For many people in Africa, livestock is the most important way of livelihood, and disease in such animals results in huge economical losses. Human and animal trypanosomiasis appears to be one of the biggest issues regarding economic development in Africa (Wilson et al., 1963). Without taking the indirect impacts of livestock on crop production, such as the use of livestock as draught animals, into account, it was estimated that the disease causes an annual loss of 1.34 billion US dollars (Kristjanson et al., 1999).

Animal reservoirs for these pathogenic African trypanosomes have long been assumed, but the importance of these reservoirs in the transmission of trypanosomiasis is not yet fully understood. In this dissertation the possibility and importance of an animal reservoir, and more specifically a wildlife reservoir, for both human and animal trypanosomiasis, is further discussed.

LITERATURE STUDY 1. The parasite: Trypanosoma Trypanosomes are flagellated, extracellular protozoa, that can cause disease in both humans and animals. They belong to the family of the Trypanosomatidae, the order of the Kinetoplastida, the phylum of Sarcomastigophora, and the subkingdom of Protozoa (WHO, 2013). Most of the pathogenic trypanosomes in Africa are transmitted by the tsetse fly. The African trypanosomes, with the exception of T. theileri, are classified in the group of Salivaria. Development of these trypanosomes takes place in the anterior part of the insect gut, and infection occurs by inoculation (Itard, 1989).

1.1. Morphology The parasite is an elongated, flat, unicellular organism, with a characteristic flagellum (Itard, 1989). The average size of an African trypanosome is 20 µm (WHO, 2013). However, the shape can change during different stages of the life cycle. For the Trypanosoma spp., the most common form is the trypomastigote, or the bloodstream form (Fig. 1). The kinetoplast is typically positioned behind the nucleus, at the posterior part of the cell. A second form is the epimastigote, which has the kinetoplast located more in the centre, and before the nucleus (Itard, 1989; Namangala and Odongo, 2014).

Fig. 1: Trypanosoma brucei spp. in a blood smear. Source: Centers for Disease Control and Prevention, United States.

Trypanosomes are morphologically distinguishable from each other. This allows for species identification, which is based on several microscopic characteristics, like size, position of the kinetoplast, and presence of a free flagellum (Uilenberg, 1998). The subspecies of Trypanosoma brucei are an exception, as a morphological difference cannot be observed. Specific molecular markers have been developed to differentiate between these species (WHO, 2013; Franco et al., 2014). The serum resistance associated (SRA) gene, which is responsible for resistance against lysis of the parasite in human serum, has been identified in T. b. rhodesiense. This gene is not expressed in T. b. brucei, T. b. gambiense, T. congolense or T. vivax (Radwanska et al., 2002; Clerinx et al., 2012). According to Uzureau et al. (2013) T. b. gambiense appears to resist trypanolytic factors in human serum through a T. b. gambiense-specific glycoprotein (TgsGP).

1.2. Life cycle The Salivarian trypanosomes are transmitted to their definitive, vertebrate hosts via hematophagous insects. This does not apply to T. equiperdum, which causes a venereal disease (Itard, 1989). Although mechanical transmission through other insects occurs, the tsetse fly (Glossina) is the only cyclical vector

of the African trypanosomes (Itard, 1989; WHO, 2013). The parasite needs the fly to develop, and achieve successful transmission. Development in the fly varies according to the species of trypanosome (Itard, 1989). In this thesis, only the life cycle of the T. brucei spp. will be discussed, as most of it is applicable to all species. Only small variations in the length of the cycle, and the path of development in the fly, exist (Vickerman et al., 1988; Roditi and Lehane, 2008; Rotureau and Van Den Abbeele, 2013).

The life cycle (Fig. 2) of T. brucei is complex, and characterised by different metabolic pathways and changes in morphology (Vickerman, 1985; Vickerman et al., 1988). The cycle begins with the feeding behaviour of the tsetse fly, which depends on blood for its nutrition. Transmission of the parasite can occur when biting the host for a blood meal, through inoculation of saliva. Saliva is injected into the host’s bloodstream to avoid coagulation and to produce vasodilation (Franco et al., 2014). When infested, the fly’s saliva contains metacyclic trypanosomes, which are the only form infective to vertebrates (Itard, 1989; Franco et al., 2014). During a bite, these trypanosomes are injected sub- dermally, and will proliferate at the site of injection. A local inflammatory response, the characteristic trypanosomal chancre, develops as result of the proliferation (Barry and Emery, 1984, as cited by Vickerman et al., 1985; WHO, 2013). The metacyclic trypanosomes will then transform into replicative, slender trypomastigotes, and non-replicative, stumpy forms. They enter the bloodstream via draining lymph nodes. The slender forms are adapted to the vertebrate host, and thus maintain the infection in the blood. Eventually, these slender trypomastigotes can penetrate the blood vessels, and excavate into connective tissue. At a later stage they can also infiltrate the central nervous system. Slender forms are capable of replicating in all body fluids.

Fig. 2: The life cycle of the Trypanosoma brucei spp. Source: Centers for Disease Control and Prevention, United States.

When an infected host is bitten by a tsetse fly, only the stumpy, non-replicative trypomastigotes in the blood meal, which are adapted to the circumstances in the insect, survive. The slender forms are rapidly

killed by proteases (Sbicego et al., 1999), or change into stumpies (Vickerman, 1985). The stumpy trypomastigotes will differentiate into procyclic forms in the midgut of the fly, after which they will continue to replicate. These procyclic trypomastigotes will then migrate to the proventriculus, where they change into longer and thinner mesocyclic trypomastigotes (Vickerman, 1985; Vickerman et al., 1988). These forms move to the salivary glands, where they change into epimastigotes, and again start multiplying. Finally, the epimastigotes will differentiate into non-replicating metacyclic trypomastigotes, which are capable of infecting the vertebrate host.

1.3. Classification Based on the life cycle and morphology, the Salivarian trypanosomes can be classified into four subgenera (Itard, 1989). A fifth subgenus (Tejeraia) can also be added, as T. rangeli was moved from the subgenus Herpetosoma (Stercoraria) to the Salivarian trypanosomes (Table 1). Like in African trypanosomes, the transmission of T. rangeli occurs through the bite of an infected insect, and not, like other Stercoraria, through contamination from the posterior end (Añez, 1982). T. rangeli can be found in Central and South America (Grisard et al., 1999), and thus will not be further discussed.

Table 1: Classification of Salivarian trypanosomes. Table based on the WHO report on human African trypanosomiasis (2013).

Subgenus Species Duttonella Trypanosoma vivax Trypanosoma uniforme Nannomonas Trypanosoma congolense Trypanosoma simiae Trypanozoon Trypanosoma equiperdum Trypanosoma evansi Trypanosoma brucei:  Trypanosoma brucei brucei  Trypanosoma brucei rhodesiense  Trypanosoma brucei gambiense Pycnomonas Trypanosoma suis Tejeraia Trypanosoma rangeli

2. The vector: Glossina All the mammalian African trypanosomes are transmitted by hematophagous insects. Like mentioned before, Tsetse flies (Glossina spp.) are the only cyclical vectors of the human and animal African trypanosomes. Mechanical transmission by Tabanidae, Stomoxyinae, and even Hippoboscidae, has also been recorded. Spread of T. evansi and T. vivax beyond the borders of the typical tsetse regions is due to these insects (Itard, 1989).

2.1. General features Tsetse flies are large, brown, but never metallic, flies, which vary in length from 6 to 16 mm. They can be recognised by the ‘hatchet’ cell on their wings (Fig. 3). Both female and male flies feed on blood, and are capable of transmitting trypanosomes. Feeding occurs every 3 to 5 days (Itard, 1989).

Fig. 3: A tsetse fly (Glossina morsitans). The ‘hatchet’ cell can be seen in the centre of the wings. Source: Illustrated lecture notes on Tropical Medicine, Institute of Tropical Medicine, Antwerp.

Glossina spp. spend the majority of their time resting on vegetation (Krinsky, 2002). Daily activity is limited to a few minutes. During the hot season, the flies are active in the early morning and late afternoon. In colder periods they are only active during the warmest hours of the day (Itard, 1989).

2.2. Distribution Tsetse flies are found solely on the continent of Africa. They are geographically restricted to 23 million km2 of sub-Saharan Africa of which 40% (9.5 million km2) is covered by tsetse infested regions (Jahnke et al., 1988). Optimal development of the fly occurs around 25 °C (Itard, 1989). They are usually not present in areas with less than 500 mm of annual precipitation (Krinsky, 2002). Extreme drought in the north (Sahara Desert) and low temperatures in the south (Namib and Kalahari Desert) thus confine their flying range to an area between the latitudes of roughly 14° N and 30° S. (Itard, 1989; Krinsky, 2002; Moloo, 1993, as cited by Franco et al., 2014). Moreover, their distribution is limited by altitudes above ca. 1500 m (Krinsky, 2002).

2.3. Taxonomy and subgenera The Glossina spp. belong to the order of Diptera (true flies) and are the only genus in the family of Glossinidae. Thirty-one species and subspecies have been described and classified into three subgenera (Annex 1) (Potts, 1973, as cited by Krinsky, 2002; WHO, 2013). This classification is mainly based on the shape of the male and female genitalia (Itard, 1989).

The subgenus Nemorhina, or the palpalis group, consist of small to medium sized tsetse flies (Itard, 1989). They are found in West and Central Africa and live in vegetation close to water, such as riverbanks and gallery forests. Flies of this subgenus are therefore referred to as ‘riverine tsetses’ (Itard 1989; WHO, 2013). Some of these species are also known to inhabit areas of agricultural activity, like coffee and cacao plantations. These plantations provide the fly with resting, breeding and feeding opportunities (Challier and Gouteux, 1980). Together with growing urbanisation, this gives rise to the tsetse’s survival in suburban and urban locations (Tongue et al., 2012).

The subgenus Glossina sensu stricto, or the morsitans group, consists of medium sized tsetse flies (Itard, 1989). They are mainly found in Central and Southeast Africa (Krinsky, 2002). They receive the name ‘savannah tsetses’ from inhabiting woodland savannahs (Itard, 1989; WHO 2013).

The subgenus Austenina, or fusca group, consists of the largest of the tsetse flies (Itard, 1989). They live in forested habitats in West and Central Africa (Krinsky, 2002), and are referred to as ‘forest tsetses’ (Itard, 1989; WHO, 2013). However, human activity is causing this subgenus to disappear (WHO, 2013).

Fig. 4: Distribution of the palpalis (A), morsitans (B) and fusca group (C). Source: Programme Against African Trypanosomes, Food and Agriculture Organization.

As seen in Fig. 4, the habitat of the Austenina (fusca) and Nemorhina (palpalis) species overlaps. The area surrounding the dense forests of equatorial Africa, provides the habitat for the Glossina s. str. (morsitans) subgenus. There are, however, several local variations in distribution. These variations depend on flora and fauna characteristics, and climate (Itard, 1989).

2.4. Feeding preferences The three subgenera prefer feeding on numerous different hosts (Krinsky, 2002). Feeding preferences are based on visual and olfactory stimuli, like color, movement, and odor of the host (Torr, 1989, as cited by Franco et al., 2014; Tirados et al., 2011). Weitz (1963) and Clausen et al. (1998) identified the

origin of vertebrate blood in the guts of, respectively 22,640 and 29,245, wild-caught Glossina species in various zones in Africa. Tables 2, 3 and 4 are based on the results of these findings.

Table 2: Feeding preferences of the Nemorhina subgenus. Based on findings of A: Weitz (1963) and B: Clausen et al. (1998).

G. palpalis G. fuscipes G. tachinoides Primates A: 38.8% A: 18.2% A: 42.7% B: 18.2% B: 8.9% B: 2.0% Suids A: 5.5% A: 3.2% A: 1.9% B: 43,8% B: 15.3% B: 0.6% Ruminants A: 22,0% A: 37.8% A: 30.4% B: 17.8% B: 22.9% B: 33.6% Other A: 4.1% A: 5.1% A: 16.0% mammals B: 7.0% B: 6.5% B: 49.7% Reptiles A: 27.7% A: 34.4% A: 8.3% B: 10.5% B: 42.1% B: 13.7%

Weitz (1963) classified G. palpalis, G. fuscipes and G. tachinoides into the group that feeds on man and most available hosts. This was largely confirmed by Clausen et al. (1998). These species are restricted to the vicinity of water and attack depends upon the extent in which the host invades the Glossina habitat. Their food sources are therefore tremendously diverse (Weitz, 1963).

Table 3: Feeding preferences of the Glossina s. str. subgenus. Based on findings of A: Weitz (1963) and B: Clausen et al. (1998).

G. morsitans G. longipalpis G. pallipides G. austeni Primates A*: 10.4% A: 1.9% A: 2.7% A: 4.9% B**: 0.7% B: 2.5% B: 2.3% B: 5.2% Suids A*: 36.4% A: 4.3% A: 29.9% A: 57.7% B**: 57.1% B: 10.2% B: 36.2% B: 89.7% Ruminants A*: 45.2% A: 91.5% A: 63.5% A: 35.6% B**: 21% B: 72.8% B: 52.2% B: 3.4% Other A*: 7.0% A: 2.2% A: 3.5% A: 1.8% mammals B**: 20.7% B: 3.9% B: 8.2% B: 0% Reptiles A*: 0.3% A: 0% A: 0.2% A: 0% B**: 0.2% B: 9.5% B: 0.6% B: 0%

*Glossina morsitans morsitans. ** No distinction between the morsitans subspecies.

Weitz (1963) classified G. longipalpis and G. pallipides into the group that feeds mainly on Bovids, G. austeni into the group that mainly feeds on Suids, and G. morsitans (spp.) into the group that feeds

on both Bovids and Suids. This was largely confirmed by Clausen et al. (1998). Other mammals, including primates (humans), also form a small part of the blood meals of this subgenus.

Table 4: Feeding preferences of the Austenina subgenus. Based on findings of A: Weitz (1963) and B: Clausen et al. (1998).

G. fusca G. fuscipleuris G. brevipalpis G. longipennis Primates A: 0% A: 0.7% A: 0.7% A: 0.4% B: 0% B: 1.4% B: 0% B: 0.25% Suids A: 13.7% A: 64.4% A: 39.4% A: 1.0% B: 7.6% B: 70.7% B: 8.6% B: 60.6% Ruminants A: 73.6% A: 19.9% A: 23.4% A: 17.9% B: 84.0% B: 21.2% B: 6.4% B: 21.5% Other A: 12.9% A: 15.0% A: 36.3% A: 73.4% mammals B: 4.2% B: 6.6% B: 85.0% B: 15.6% Reptiles A: 0% A: 0% A: 0% A: 0% B: 3.4% B: 0% B: 0% B: 0.25%

Weitz (1963) classified G. fusca into the group that feeds mainly on Bovids, G. fuscipleuris into the group that mainly feeds on Suids, and G. longipennis and G. brevipalpis into the group that feeds mainly on other mammals. With the exception of G. longipennis and G. brevipalpis, who’s feeding preferences differ from the 1963 report, this was also largely confirmed by Clausen et al. (1998).

The feeding preferences of each Glossina species seem to be characteristic and not fully dependent on the availability of hosts (with the exception of the palpalis group). This suggests a genetic background, and is supported by the fact that some common wild animals, such as zebra, oryx, and wildebeest, are almost never bitten (Weitz, 1963; Franco et al., 2014). It is assumed that their colours are less compelling to the fly (WHO, 2013), or that their skin contains repellent substances (Saini and Hassanali, 2007). Tsetse flies also rarely feed on birds. An exception to this is G. longipennis. Of its 1422 blood meals 7.3% could be traced back to bird origin, mainly ostrich (Weitz, 1963).

2.5. Important vector species All Glossina species are capable of transmitting African trypanosomes, though only a few are important in spreading human and animal trypanosomiasis (WHO, 2013; Franco et al., 2014).

The riverine tsetses (subgenus Nemorhina) are the most important vectors of human African trypanosomiasis (WHO, 2013). They seem to be attracted to man, a trait that is not frequently displayed in other Glossina species (Weitz, 1963). G. palpalis, but also G. tachinoides (Krinsky, 2002), appears to be responsible for transmitting T. b. gambiense in West Africa. This in contrast with G.fuscipes, which spreads both T. b. gambiense and T. b. rhodesiense throughout Central and East Africa (Krinsky, 2002; WHO, 2013). G. fuscipes and G. palpalis are also vectors of animal trypanosomes (WHO, 2013).

The distribution of the Glossina s. str. subgenus is related to the presence of wild fauna and cattle (WHO, 2013; Franco et al., 2014). G. pallidipes, G. swynnertoni (not included in the tables), and G. morsitans spp. are responsible for the transmission of T. b. rhodesiense in East Africa (Molyneux and Ashford, 1983, Krinsky, 2002; WHO, 2013). G. swynnertoni is also a significant vector for animal trypanosomes (WHO, 2013). This species belongs to the group that feeds mainly (65,4%) on Suids (Weitz, 1963).

The Austenina species have not been known to transmit sleeping sickness, but they are effective vectors of animal trypanosomes. However, they often live far away from grazing grounds, and are thus less important in transmitting disease in domestic animals (WHO, 2013; Franco et al., 2014).

3. Human African trypanosomiasis Human African trypanosomiasis (HAT), or sleeping sickness, is a disease affecting people in rural settings of sub-Saharan Africa (Brun et al., 2009). The interaction between the human host, the Glossina spp. and the trypanosomes is very complex, resulting in a focal geographical distribution of HAT. An estimated 57 million people in Africa, distributed over 1.38 million km2, are at risk of contracting sleeping sickness caused by T. b. gambiense. Approximately 12.3 million people in a region of 0.171 million km2 are at risk of getting infected with T. b. rhodesiense (WHO, 2013).

3.1. Causative agents Sleeping sickness is caused by T. b. gambiense or T. b. rhodesiense. The latter causes an acute disease in East and Southern Africa, while T. b. gambiense is responsible for a more chronic form in Central and West Africa (Molyneux and Ashford, 1983; Itard, 1989; Brun et al., 2009; WHO, 2013). In Uganda, some regions have already shown overlap between T. b. gambiense en T. b. rhodesiense. There is a possibility that these two forms will merge completely in the future (Picozzi et al., 2005; Berrang-Ford et al., 2006). T. b. gambiense is responsible for 97% of the HAT cases in the last decade (Simarrro et. al., 2011), whereas T. b. rhodesiense seems to only accidentally infect humans (Franco et al., 2014).

3.2. Clinical symptoms The disease can be divided into an initial haemo-lymphatic phase and a subsequent meningo- encephalitic phase, where the central nervous system is invaded by trypanosomes (Blum et al., 2005). Symptoms of these stages can overlap, making it hard to distinguish between them (Kennedy, 2013).

3.2.1. Clinical symptoms of T. b. gambiense HAT caused by T. b. gambiense has an average duration of 3 years, with the two stages evenly divided between the length of the disease (Checchi et al., 2008). A trypanosomal chancre (Fig. 5) is very rarely observed in these cases (Malvy and Chappuis, 2011; Brun and Blum, 2012). Chronic, intermittent fever, headache, lymphadenopathy, pruritis, anaemia and weakness are some of the most common symptoms of the first stage. Swelling of the posterior cervical lymph nodes (Winterbottom’s sign; Fig. 5) is more typical for gambiense sleeping sickness (Kennedy, 2013; WHO, 2013). Some endemic HAT areas overlap with filariasis regions, which means the pruritis could also be explained by the presence of these parasites (Blum et al., 2005). Hepatosplenomegaly is frequently seen, and even cardiac problems are possible, although heart failure is not often reported (Blum et al., 2007). Oedema of the face and deep hyperaesthesia (Kerandel’s sign) have also been observed (Malvy and Chapuis, 2011).

In the second stage of the disease, sleep disturbances and neurological disorders dominate. According to Buguet et al. (2004) the disease causes dysregulation of the sleep-awake cycle, and fragmentation of the sleep pattern, rather than the inversion of sleep that is normally reported for HAT. Blum et al. (2005) studied 2541 patients with trypanosomiasis, of which 74% showed the typical sleep disorders that give sleeping sickness its name. Headache, mood and behavioural changes are commonly found due to the meningo-encephalitis caused by the trypanosomes invading the central nervous system. Motor weakness, abnormal movements, tremor, walking difficulties, problems with speech, and even psychiatric disorders, like depression and delirium, have been witnessed during this phase (Blum et al.,

2005; Kennedy, 2006). In the terminal stage, the cachectic patient develops incontinence, cerebral oedema and advanced mental impairment, which ultimately leads to death (Kennedy, 2006; Malvy and Chappuis, 2011). However, the clinical features of sleeping sickness can be highly variable between individuals and foci (Blum et. al, 2005; Kennedy, 2013).

Fig. 5: Left: A trypanosomal inoculation chancre. Right: Winterbottom’s sign. Source: Illustrated lecture notes on Tropical Medicine, Institute of Tropical Medicine, Antwerp.

3.2.2. Clinical symptoms of T. b. rhodesiense Rhodesiense HAT is more acute, continuing to the second stage within weeks and leading to death within 6 months (Odiit et al., 1997). Most symptoms are similar to T. b. gambiense infection, but trypanosomal chancres and oedema are seen more often (Malvy and Chappuis, 2011; Brun and Blum, 2012; WHO, 2013). Swelling of the lymph nodes tends to be located in the submandibular, axillary or inguinal region, instead of posterior cervical, like in the gambiense form (WHO, 2013).

3.2.3. Clinical symptoms in non-native individuals Cases have also been recorded outside of Africa, mostly in travellers that return from visits to game parks (Jelinek et al., 2002; Urech et al., 2011; Clerinx et al., 2012). Symptoms in patients from non- endemic countries are different from the symptoms in African people suffering from HAT. Disease due to T. b. rhodesiense is mostly seen in travellers, whereas T. b. gambiense infections are rare in travellers, and occur more in immigrants (Blum et al., 2011). The onset of both diseases is more rapid in non-native patients, which leaves no room for the classic neurological signs and sleep disturbances to be developed. Chancres were seen more often in both infections. Symptoms like nausea, vomiting and diarrhoea were also reported in T. b. rhodesiense patients (Blum et al., 2011; Urech et al., 2011).

4. Animal African trypanosomiasis Together with HAT, animal trypanosomiasis is a major cause of rural underdevelopment in sub-Saharan Africa (Brun et al., 2009). Unlike HAT, animal trypanosomiasis is much more widespread. It is a major constraint to livestock production in 40 sub-Saharan African countries. About 50 million cattle and 70 million small ruminants are annually at risk of contracting the disease (Coustou et al, 2012).

3.1. Causative agents The diseases caused by pathogenic trypanosomes in domestic animals are respectively known as nagana, surra and dourine (Molyneux and Ashford, 1983; Itard, 1989; Namangala and Odongo, 2014; Maes, 2014). Surra is a disease caused by T. evansi and mainly affects camels, while dourine, caused by T. equiperdum, is a sexually transmitted disease in horses. Nagana is the best known disease complex, as it affects several domestic animals and is caused by several trypanosome species. However, it is most important in cattle and small ruminants, as they are the most frequently reared animals in sub-Saharan Africa (Namangala and Odongo, 2014). Nagana can cause significant economic losses in livestock due to infection with T. congolense, T. vivax and to lesser extent with T. b. brucei (Losos and Ikede, 1972; Clarkson, 1976; Molyneux and Ashford, 1983). The disease caused by T. vivax is sometimes also referred to as souma (Maes, 2014).

T. congolense is accountable for more than 80% of the AAT cases in domestic animals in West, Central and Southern Africa (Simukoko et al., 2007). Tsetse flies of the Nemorhina subgenus seem to be less susceptible to infection with this species than the other groups of tsetse flies (Molyneux and Ashford, 1983). T. vivax is de second most important trypanosome to cause nagana, and infection results in a milder form of disease than T. congolense (Namangala and Odongo, 2014). It accounts for almost all the AAT cases in West-Africa (Adam et al., 2012) and is most commonly found in Bovids (Clarkson, 1976; Molyneux and Ashford, 1983). T. vivax can be transmitted by all Glossina species (Molyneux and Ashford, 1983) and is often found outside the tsetse belt due to mechanical transmission via other insects (Itard, 1989). T. b. brucei is the widest spread African trypanosome, and can infect many species of domesticated and wild animals (Clarkson, 1976). However, it has a relatively low pathogenicity in ruminants (Namangala and Odongo, 2014). Tsetse flies of the Austenina group have not been known to transmit T. brucei spp. (Krinsky, 2002, WHO, 2013). Simultaneous infection with one or more of these species is not uncommon (Molyneux and Ashford, 1983; Eshetu and Begejo, 2015).

4.2. Clinical symptoms The pathology of animal trypanosomiasis differs within each host and each parasite species (Losos and Ikede, 1972). The general symptoms, however, are often very similar (Molyneux and Ashford, 1983). A trypanosomal chancre is hardly ever seen in natural infections. The most dominant, pathogenic feature in cattle is anaemia (Molyneux and Ashford, 1983; Van den Bossche and Rowlands, 2000). Other symptoms include intermittent fever, lymphadenopathy, lacrimation, weakness, lethargy, weight loss and sometimes oedema. In the chronic form of the disease (Fig. 6), neurological signs, like weakness of the hind limbs, and sometimes even paresis or paralysis, can be seen. Eventually, the animal will die of cachexia (Losos and Ikede, 1972, Itard, 1989; Krinsky, 2002; Namangala and Odongo, 2014). Most

organ systems are infected and pathological lesions like myocarditis, lung oedema and hepatosplenomegaly are often seen. However, the infection can also cause acute disease, as not all animals will survive the high fever and severe haemolytic first phase of the disease (Losos and Ikede, 1972; Itard, 1989; Molyneux and Ashford, 1983; Eshetu and Begejo, 2015). Sudden death is more seen in animals that recently have been introduced in tsetse infested areas, whereas the chronic form appears more in endemic areas (Namangala and Odongo, 2014). Widespread visceral and mucosal haemorrhaging has also been reported among cattle infected with T. vivax (Molyneux and Ashford, 1983). The word ‘nagana’ is Zulu for ‘being in depressed spirit’, which is the state of the animal in the terminal stage of the disease (McKelvey, 1973, as cited by Krinsky, 2002). These clinical signs and lesions are not fully diagnostic for trypanosomiasis, as several other conditions, like piroplasmosis, can also cause these symptoms (Losos and Ikede, 1972).

Fig. 6.: Chronic form of a trypanosome infection in cattle. Source: International Livestock Research Institute, Kenya.

Because of the fever associated with the disease, abortion is a frequent symptom among pregnant animals. Male infertility due to testicular damage is also reported (Molyneux, 1983; Eshetu and Begejo, 2015). Cattle kept in areas of AAT seem to have lower calving rates, lower milk yields and higher rates of calf mortality. Animals with chronic infection are often also too weak to be used as draught animals, which has an indirect impact on crop production (Swallow, 1999).

Some native breeds of cattle, like N’dama, are capable of tolerating trypanosomes without falling seriously ill or without having considerable production losses. This phenomenon is known as ‘trypanotolerance’. Zebu cattle, on the other hand, appear to be more susceptible for trypanosomiasis (Murray et al.,1981; Paling et al., 1991 as cited by Naessens, 2005). The use of trypanotolerant breeds is currently exploited to reduce effects of animal trypanosomiasis on livestock and crop production (Molyneux and Ashford, 1983; Krinsky, 2002; Eshetu and Begejo, 2015).

5. Reservoirs Salivarian trypanosomes have an extremely wide host range (Molyneux and Ashford, 1983). Ruminants, pigs and carnivores, both domestic and wild, all seem to be susceptible to T. brucei spp. (Itard, 1989). Although it was already shown in previous studies (Mehlitz, 1982, as cited by Njiokou et al., 2005) that wild and domestic animals could act as a reservoir to HAT, early identification methods for differentiating between trypanosomes lacked in sensitivity and specificity. With the development of PCR, identification of different trypanosomes within vertebrate hosts has been significantly improved (Biteau et al., 1999).

5.1. Reservoir for human African trypanosomiasis 5.1.1. Reservoir of T. b. gambiense Control measurements directed at the human reservoir of sleeping sickness have been successful in reducing its transmission. This suggests that the infection with T. b. gambiense is sustained by a man- fly-man cycle. The 3-year-long duration of the disease also supports this perception (Molyneux and Ashford, 1983; WHO, 2013). Despite the general acceptance of a human reservoir, elimination of the disease in certain T. b. gambiense foci could not always be achieved by surveillance and control. It is reported that this might be due to under-detection of cases during human population screenings (Checchi et al., 2012). However, animal reservoirs for T. b. gambiense have also been suggested, but still need to be clearly identified (Molyneux and Ashford, 1983; WHO, 2013).

Simo et al. (2006) examined 133 blood samples of pigs from Fontem, a sleeping sickness focus in Cameroon, to investigate the role of a possible animal reservoir. Through PCR, they found a high prevalence (73.7%) of trypanosomes in the samples. Of the infected pigs, 40% was infected with T. brucei spp., and 15.8% of these were infected with T. b. gambiense. Although high in parasitaemia, symptoms of trypanosomiasis were not seen. This is consistent with Itard (1989), who noted that T. brucei spp. are not very pathogenic in pigs. It was concluded that the pigs from the Fontem focus probably played an important role as (asymptomatic) reservoir species for sleeping sickness.

A larger screening for T. b. gambiense was done in the Mbini and Kogo regions of Equatorial Guinea (Cordon-Obras et al., 2009). Of the 698 animals (456 goats, 218 sheep and 24 pigs) that were sampled, 39.5% was infected with trypanosomes. In the Mbini region 52.6% of the 346 animals sampled were positive for T. brucei spp., whereas 36.1% of the 352 animals in Kogo were infected. PCR showed that only 2% of the animals (goat and sheep, no pigs) from Mbini were infested with T. b. gambiense. In Kogo none of the animals were tested positive for T. b. gambiense. It was implicated that these results raised a possibility for an animal reservoir, but to confirm this further studies were needed. The authors also noted that more circulation of sheep and goat would result in a greater risk of infection due to the possibility of the G. palpalis to adapt to, and thus infect, every host available.

Njiokou et al. (2010) investigated the prevalence of T. b. gambiense in four domestic animal species (sheep, goat, pigs and dogs), found in four different active HAT foci in Cameroon. T. brucei spp. were detected in 19.88% of the 875 (307 pigs, 267 sheep, 264 goats and 37 dogs) animals sampled. With PCR it was seen that 3.08% out of the total sampled animals had DNA specific for T. b. gambiense. Sheep were most infected, followed by goats and pigs. T. b. gambiense infection in dogs was not

detected with PCR. Again, the authors noted that, although it was obvious that these animals will possibly have a role in HAT transmission, further research needs to be done to clarify their importance.

Lack of a domestic animal reservoir has also been reported by Balyeidhusa et al. (2011). In Uganda, the only country that has been affected by both T. b. gambiense and T. b. rhodesiense (WHO, 2013), a total of 3267 bloodsamples were taken from domestic animals. Although 12.8% of these blood samples were tested positive on infection with Trypanozoon species, none of these harboured T. b. gambiense. It was concluded that even though domestic animals are susceptible to infection with T. brucei spp., none of the investigated domestic animals in Uganda were infected with T. b. gambiense.

Wild animals have also been reported to be infected by T. b. gambiense. Herder et al. (2002) sampled 164 animals (54 primates, 45 ungulates, 39 rodents, 11 carnivores, 10 pangolins and 5 reptiles) in the forest belt in Cameroon. Of these 164 animals 8% was carrying T. brucei gambiense. The parasite was found in the brush-tailed porcupine, giant rat, black striped duiker, blue duiker, white-eyelid mangabey, greater white-nosed monkey, palm civet and small-spotted genet. T. b. gambiense had never before been identified in palm civets or small-spotted genets. However, the presence of the parasite in these blood samples, like in the samples from the domestic animals, does not specify the importance of that animal as a reservoir. To indicate the significance of these animals as reservoirs, a larger number of wild fauna was sampled by Njiokou et al. (2005) in Cameroon. During the survey, 1142 animals (253 primates, 234 ungulates, 237 rodents, 49 pangolins, 45 carnivores, 9 reptiles and 5 hyraxes) were sampled from four different regions. Of these, three regions were known HAT foci, and one, in which sleeping sickness was never reported, was used as a control zone. The animals that were noted to be carriers in the report of Herder et al. (2002) were confirmed as carriers of T. b. gambiense in this study. T. b. gambiense was detected in 1.6% of the animals, but the parasite was not found in wild animals in the control zone. Both these studies confirmed a potential role of wild fauna in the persistence of HAT.

However, the results of the studies above are not conclusive, as the T. b. gambiense strains found in wild and domestic animals are not per se infective to humans (Njiokou et al. 2005; Simo et al., 2006).

Research was also conducted on wild non-human primates (chimpanzees i.a.). Although it was established that chimpanzees are often infected with T. brucei spp., attempts to distinguish between the subspecies were futile (Jirku et al., 2015). It was noted in Godrey and Killick-Kendrick (1967) that T. b. gambiense did not cause noticeable symptoms in these primates, despite presence of the trypanosome in the cerebro-spinal fluid. Chimpanzees might therefore act as reservoir hosts for T. b. gambiense.

Another hypothesis for persistent HAT transmission is the existence of asymptomatic carriers, or seropositive individuals with negative parasitology. The parasitaemia in these patients is so low that tests are unable to detect it. This trypanotolerance presents a possibility for a long lasting human reservoir that causes persistence of HAT in some foci (Koffi et al., 2006; WHO, 2013).

5.1.2. Reservoir of T. b. rhodesiense

It has been known for quite some time that T. b. rhodesiense is a zoonotic disease and that the parasite maintains its population through an animal reservoir (Heisch et al., 1958; WHO, 2013). G. pallipides, G.

morsitans, G. swynnertoni and G. fuscipes, which were identified as carriers of T. b. rhodesiense (WHO, 2013), feed on a wide variety of wild and domestic animals (Weitz, 1963; Clausen et al., 1998). This indicates the complexity of the transmission cycle of T. b. rhodesiense.

In a study by Onyango et al. (1966), the objective was to find out whether domestic animals, like cattle, were carrying human-infective trypanosomes. Blood samples were taken from 203 Zebu cattle. A total of 68 animals were found positive of infection with trypanosomes. Of these 68 positive samples, 43 isolates could be obtained. On blood films was seen that all these isolates were polymorphic trypanosomes, which is typical for trypanosomes of the T. brucei spp. Two different isolates were then inoculated in two human volunteers. The first volunteer (A) was inoculated with an EATRO 835 isolate, whereas the second volunteer (B) was injected with an EATRO 839 strain. In volunteer A pain and swelling at the inoculation site were witnessed almost directly. Symptoms like fever, headache, pain at the back of the neck, and swelling of lymph nodes were also noted. Trypanosomes were seen in a blood film on day 9 after inoculation. On day 12 the patient was better, but in order to ensure infection with T. b. rhodesiense, treatment was put on hold until a second wave of clinical symptoms was noticed. This was due to the fact that these symptoms could have been caused by a transient parasitaemia of a trypanosome non-infective to man. Patient A was treated after a second wave of parasitemia was witnessed. In volunteer B the inoculation didn’t cause any clinical symptoms, apart from a small swelling at the injection site. Blood films were negative for trypanosomes throughout the length of the study.

Volunteer B was later used in another study with EATRO 835 (Van Hoeve et al., 1967) to investigate the cyclical transmission of this strain to humans, cattle and sheep. After 10 rats were inoculated with this T. b. rhodesiense isolate, 937 G. morsitans flies had the chance to feed of these infected rats. Two sheep were then offered to 680 of these infected tsetse flies, and subsequently became infected. One infected G. morsitans was then offered a blood meal on a Zebu cow and after 6 days the first trypanosomes were detected in a blood smear. The cow was left untreated to indicate the duration of the infection and thereby the time the cow could serve as a possible reservoir for T. b. rhodesiense. The cow was positive until the 245th day of the infection. Eight infected G. morsitans were then fed on volunteer B from the study done by Onyango et al. (1966). The same clinical symptoms that volunteer A showed in the first study, were also witnessed on volunteer B. The blood films were however positive from the beginning of the treatment, so treatment began immediately. It wasn’t necessary to confirm the pathogenicity of strain, as the chance of a transient parasitaemia was rather small.

It was concluded from the study of Onyango et al. (1966) that rhodesiense HAT could be mechanically transferred from cattle to man. The second study (Van Hoeve et al., 1967) proved that cyclical transmission of this strain to humans, cattle and sheep was also possible. Although a trypanosusceptible species, the Zebu remained in good shape. Like T. b. brucei, T. b. rhodesiense is not pathogenic to cattle (Fèvre et al., 2001; Namangala and Odongo, 2014). Such cattle would therefore not be presented for treatment, and these animals could thus continue to be a potential reservoir for T. b. rhodesiense. he authors recommended mass treatment of livestock during outbreaks of rhodesiense HAT.

According to Hide et al. (1996) tsetse flies were five times more likely to pick up an T. b. rhodesiense infection from cattle than from humans.

Fèvre et al. (2001) investigated the cause of an outbreak of rhodesiense HAT in the Soroti region in Uganda, where sleeping sickness due to this trypanosome was never recorded before. It was suspected that the outbreak was caused by the import of cattle from markets in endemic HAT areas. A good 54% (1510 animals) of 2796 cattle that were traded at the main cattle market in the Soroti district, came from T. b. rhodesiense foci. Distance from the cattle market also seemed a significant risk factor to humans for contracting sleeping sickness. These findings suggested an association between the outbreak and the movement of cattle from endemic HAT areas. Like Oyangu et al. (1966) and Van Hoeve et al. (1967) the authors also suggested that treatment of cattle should be necessary to prevent import of rhodesiense sleeping sickness into new areas.

In the Totoro and Soroti regions in Uganda samples were also taken by Wellburn et al. (2001). Of the 41 cattle blood samples that were collected in Totoro, the SRA gene for T.b. rhodesiense was present in eight cases. Another 200 cattle were sampled in the Soroti district, and a prevalence of 18% for T. b. rhodesiense infection was found. It was concluded that this proved the central role of cattle in the transmission and persistence of T. b. rhodesiense HAT in these regions.

The existence of a wildlife reservoir for T. b. rhodesiense was firstly proven by Heisch et al. (1958). Blood samples were taken from 24 animals (13 duikers, 10 bushbucks and one serval cat) in the Nyanza region, Kenya. Strains of polymorphic T. brucei spp. were isolated from both the duiker and the bushbuck. The isolates were inoculated into human volunteers and the strain from the bushbuck appeared to pathogenic to man. This outcome established that T. b. rhodesiense had zoonotic potential.

Knowledge on wildlife as a reservoir for T. b. rhodesiense (and other trypanosomes, like T. congolense) has been improved since then. However, it seems difficult to collect a sufficient amount of samples from different wildlife species to conduct a good epidemiological analysis (Anderson et. al., 2011).

A survey in and around the Serengeti National Parak, Tanzania, was carried out on 95 animals (31 spotted hyenas, 43 lions, 20 hartebeests and one waterbuck) by Geigy and Kauffmann (1973). Blood samples were collected from the immobilized animals and tested for trypanosomes. A total of 74 animals were found to be infected, 28 (10 hyenas, 15 lions, three hartebeests) of which were infected with T. brucei spp. According to Bertram (1973) 7.5% of the large mammals in Serengeti National Park, Tanzania, are infected with T. brucei spp. The great majority of these appeared to be wildebeest, due to their enormous abundance. Although Geigy and Kauffmann (1973) recorded that lions and hyenas could be important as reservoirs due to their high infection rate with T. brucei spp., Bertram (1973) concluded that lions and hyenas appeared to be less important as reservoirs, compared with more abundant, less infected species.

In Tanzania, Kaare et al. (2006) completed a study to determine the prevalence of T. b. rhodesiense and other trypanosomes in livestock and wildlife in and near the Serengeti National Park, Tanzania. Blood samples were taken and PCR was done to identify all species. In 220 wild animals (68 wildebeests, 46 topis, 26 zebras, 24 Thompson’s gazelles, 21 warthogs, 15 impalas, nine lions, four elands, two spotted hyenas, one cheetah, one buffalo, one giraffe, one oribi and one reedbuck), T. brucei spp. was found in the spotted hyena, lion, reedbuck, topi, warthog and wildebeest. Warthogs

showed the highest prevalence of T. b. rhodesiense at 9.5%. A prevalence of 1.1% of T. b. rhodesiense was found in the 518 cattle sampled in that region. It was concluded that control of sleeping sickness in such an area may also depend on limited interaction between wildlife and livestock.

Anderson et al. (2011) did a survey on trypanosome prevalence in 418 wildlife species in the Luangwa Valley, Zambia. In this area, large populations of tsetse flies and abundance of wildlife can be found, while livestock keeping is almost non-existent. The prevalence of trypanosome infection in these species was 13.9%. Of these trypanosomes, 5.7% belonged to T. brucei spp. The majority of the T. brucei spp. infections were detected in four species: bushbuck, leopard, lion and waterbuck. Two T. b. rhodesiense positive samples were found, one in a busbuck and one in a buffalo. This was 8.3% of the T. brucei spp. that were found. In this area, It was noted that the prevalence of T. b. rhodesiense could be underestimated. Detection of T. b. rhodesiense is done on the SRA gene. However, this is only a single copy gene, and although primers amplifying another gene were included as a positive control, failure to detect this other gene, might result in failure to detect SRA positive samples. It was also assumed that cross-immunity due to infection with a genetically diverse species, such as T. congolense, led to partial protection of these animals against infection with T. b. rhodesiense. As it was the first time a T. b. rhodesiense was detected in a buffalo, and a T. brucei spp. was detected in a leopard, it was concluded that the reservoir appeared to be more widespread than previously mentioned.

5.2. Reservoir for animal African trypanosomiasis Apart from being suitable hosts for human sleeping sickness, numerous wild animal species also seem to be naturally infected with T. vivax, T. congolense and T. b. brucei (Mulla and Rickman, 1988). Certain species, like gazelle, dik-dik, jackal, bat-eared fox and aardvark, seem to usually die as a result of the infection, whereas other species, like eland, hyena, bushbuck and impala are susceptible to infection and remain parasitaemic for quite a while. Warthogs, bush pigs and porcupines only seem to show transient infection (Ashcroft et al., 1959, as cited by Mulla and Rickman, 1988).

Mulla and Rickman (1988) also recorded that the level of parasitaemia and anaemia in wildlife was much lower than in domestic animals. It was suggested that the level of infection in wild animals can be controlled by specific host anti-bodies, efficient phagocytosis, non-immunological responses and innate trypanolytic factors. This phenomenon is called ‘trypanotolerance’ and is an important aspect of their role as a wildlife reservoir for both human and animal trypanosomiasis.

A survey in and around the Serengeti National Park, Tanzania, was carried out on 95 animals (31 spotted hyenas, 43 lions, 20 hartebeests and one waterbuck) by Geigy and Kauffmann (1973). Blood samples were collected from the immobilized animals and tested for trypanosomes. A total of 74 animals were found to be infected, 23 of which harboured T. congolense (7 in hyenas, 15 lions and one hartebeest). Two cases of T. vivax were reported in hartebeest. According to a report of Bertram (1973) lions were found to carry either T. brucei spp. or T. congolense. Their infection rate is amongst the highest in animals, though they rarely get bitten by Glossina spp. This seemed to be consistent with the findings of Clausen et al. (1998), who reported only 3 blood samples (in G. pallipides), from a total of 13,145 samples that were identifiable up to species level, to be of lion origin. It is assumed that lions

become infected from their prey, through lesions in the oral mucosa (Molyneux and Ashford, 1983; Baker, 1968, as cited by Anderson et al., 2011). It might also be possible for lions to get infected with trypanosomes while grooming each other (Bertram, 1973).

Identification of T. congolense and T. vivax in wildlife in the Serengeti National Park was also carried out by Kaare et al. (2006). T. congolense was identified in buffalo, eland, giraffe, impala, lion, reedbuck, Thompson’s gazelle, topi, warthog and wildebeest, indicating a very large potential reservoir for this trypanosome. T. vivax was found only in warthogs and zebra.

The same was done Anderson et al (2011) in the Luangwa Valley in Zambia. The overall prevalence of T. congolense in a total of 418 wildlife samples was 6.0%, while T. vivax only had a prevalence of 3.1%. The prevalence of T. congolense was highest in the greater kudu and lion, but bushbuck, wildebeest, warthog, puku, impala and buffalo were also found to be infected. Reedbuck, waterbuck, warthog, buffalo and hippopotamus were infected with T. vivax. It was concluded that the wildlife reservoir of T. congolense would appear to be larger than the reservoir for other trypanosome species. Members of the Bovidae family seemed most frequently represented as reservoirs. Two transmission routes could be followed for T. congolense. One involving ungulate species, on which tsetses often feed, and one involving carnivores with oral transmission. The authors noted such conclusions couldn’t be made for T. vivax, as the prevalence of the trypanosome in wildlife was much lower. However, it is likely that that the Bovinae subfamily plays an important role as reservoir in the transmission of T. vivax. In the recent years an influx of people and livestock occurred in several districts of the Luangwa valley, which has led to new wildlife-livestock-human interactions. A survey of the prevalence of trypanosomes in domestic livestock in one of the regions, showed a prevalence of 28.4% in cattle. T. congolense was identified in 82.4% and T. vivax in 24.5% of the infected cattle (Mubanga, 2008, as cited by Anderson et al., 2011).

The extent of the trypanosome diversity in wildlife in the Serengeti National Park, Tanzania, and the Luangwa Valley in Zambia, was established by Auty et al. (2012). A large number of trypanosome species was identified, including species, like T. godfreyi, that were not identified in wildlife before. T. godfreyi was previously isolated from tsetse flies (McNamara et al., 1994, as cited by Auty et al., 2012) and when experimentally infected in domestic pigs, it caused a chronic disease. The T. vivax that was found in a buffalo was matched with T. vivax from a cow in Kenya, while sequences from other animals appeared to differ from the T. vivax strain in both of them. Because of their shared phylogeny, buffalo and cattle may be more likely to be susceptible to similar pathogenic strains. However, the author concluded that more information on the circulation of different strains in and between wildlife and livestock is needed to confirm such a hypothesis.

5.3. Control of reservoirs Treatment and control of the animal reservoir for T. b. gambiense is not carried out, as humans are considered to be the most important reservoir species. However, T. b. rhodesiense is a zoonosis and control of the animal reservoir for that disease is considered to be very important (WHO, 2013). A way of controlling T. rhodesiense in reservoir animal hosts is by using insecticide pour-ons in cattle. Resistance to these chemotherapeutical products is reported in animal trypanosomes (Geerts et al.,

2001), but there is no evidence to suspect resistance against T. b. rhodesiense. Vector control, which aims at reducing the population of tsetse flies, is a widely used method to control the transmission of both human and animal trypanosomiasis. According to the WHO (2013), direct control of trypanosomiasis in wildlife is not an option. Many wild animals are protected species and mass screenings would be unethical and too expensive. Game elimination, as a way to control the wildlife reservoir, was done in the past (Clarke, 1964), but these methods are no longer used because of their negative influence on biodiversity. Avoidance of wildlife areas, personal protective measurements and vector control are methods that might be useful in reducing transmission via wildlife.

DISCUSSION Although much research is done to understand the interactions between wildlife, domestic animals and humans in the transmission cycle of the African trypanosomes, it remains a complex topic. The complexity can be seen in every aspect of the transmission cycle. Although African trypanosomes are only cyclically transmitted by tsetse flies, their habitat preferences have ensured the presence of these pathogens in many African countries. The thirty-one species and subspecies of Glossina are found in abundance in forests, savannahs and watery environments in sub-Saharan Africa, where they can come in contact with multiple species of wildlife, but also humans and domestic animals. Some subspecies even inhabit peri-domestic areas and regions with agricultural activity, which allows for even more interaction between these insects and possible human hosts. As seen from the feeding preferences, tsetse flies feed on a wide variety of hosts. Feeding preferences of some of the subspecies, mostly the Nemorhina group, seemed not always based on an intrinsic, genetic background. Although specific subspecies were indicated as carriers of animal and human trypanosomes, all Glossina spp. are able to transmit trypanosomes, which complicates the epidemiology of the disease even more.

While human African trypanosomiasis is only caused by two subspecies of T. brucei, disease in animals can be caused by a much wider variety of trypanosomes, each with different transmission cycles and pathogenicity. Control measurements and surveillance systems seem to be better adapted to detect and treat human cases than to identify animal trypanosomiasis. This suggests the problem of a domestic animal reservoir for other (domestic) animals, but definitely also for humans. Asymptomatic human carriers can however also be a cause of the persistence of HAT in some areas. Although there are initiatives to eliminate HAT, the presence of such an animal reservoir and asymptomatic human carriers can be an obstacle in eradicating the disease.

A wildlife reservoir for trypanosomes infective to both domestic animal and humans has been described multiple times, although the importance of such reservoirs in the epidemiology of AAT and HAT still remains somewhat vague. However, the abundance of competent wildlife hosts in specific HAT and AAT regions throughout sub-Saharan Africa, is still a reason to suspect an important connection. As eliminating all wild animals from tsetse infected areas, to control the disease, is in conflict with conservation efforts, other solutions for the growing wildlife-livestock-human interface need to be found. Not only can migrating wildlife spread the disease from endemic to non-endemic areas, also human migrants can play a role in the emergence of new disease areas. Migration of people and domestic animals from non-endemic to endemic areas, or migration from endemic to non-endemic areas, can also ensure new outbreaks of human and animal trypanosomiasis.

Many authors noted in their articles that several perceptions on trypanosome transmission need more research in order to solve certain questions regarding wildlife reservoirs. Furthermore, there are numerous other aspects to trypanosomes and their transmission, that have not been discussed in this dissertation. That makes it difficult to make one good conclusion about the importance of wildlife reservoirs in the spreading of trypanosomes. All in all, the epidemiology of human and animal African trypanosomiasis remains complex and not quite clear. More research on the importance of wildlife as a reservoir is needed to get a better understanding of the epidemiology of the African trypanosomiases.

REFERENCES 1. Adam Y., Marcotty T., Cecchi G., Mahama C.L., Solano P., Bengaly Z., Van den Bossche P. (2012). Bovine trypanosomosis in the Upper West region of Ghana: Entomological, parasitological and serological cross-sectional surveys. Research in Veterinary Science 92, 462-468. 2. Anderson N.E., Mubanga J., Fèvre E.M., Picozzi K., Eisler M.C., Thomas R., Welburn S.C. (2011). Characterisation of the wildlife reservoir community for human and animal trypanosomiasis in the Luangwa Valley, Zambia. PLoS Neglected Tropical Diseases 5, e1211. 3. Añez N. (1982). Studies on Trypanosoma rangeli Tejera, 1920 IV – A reconsideration of its systematic position. Memórias do Instituto Oswaldo Cruz 77, 405-415. 4. Auty H., Anderson N.E., Picozzi K., Lembo T., Mubanga J. Hoare R., Fyumagwa R.D., Mable B., Hamill L., Cleaveland S., Welburn S.C. (2012). Trypanosome diversity in wildlife species from the Serengeti and Luangwa Valley ecosystems. PLoS Neglected Tropical Diseases 6, e1828. 5. Balyeidhusa A.S.P., Kironde F.A.S., Enyaru J.C.K. (2011). Apparent lack of a domestic animal reservoir in gambiense sleeping sickness in northwest Uganda. Veterinary Parasitology 187, 157-167. 6. Berrang-Ford L., Odiit M., Maiso F., Waltner-Toews D., McDermott J. (2006). Sleeping sickness in Uganda: Revisiting current and historical distributions. African Health Sciences 6, 223-231. 7. Bertram B.C.R. (1973). Sleeping sickness survey in the Serengeti Area (Tanzania) 1971. III. Discussion of the relevance of the trypanosome survey to the biology of large mammals in the Serengeti. Acta Tropica 30, 36-48. 8. Biteau, N., Bringaud F., Gibson W., Truc P., Baltz, T. (1999). Characterization of Trypanozoon isolates using a repeated coding sequence and microsatellite markers. Molecular and Biochemical Parasitology 105, 185-201. 9. Blum J.A., Burri C., Hatz C., Kazumba L., Mangoni P., Zellweger M.J. (2007). Sleeping hearts: The role of the heart in sleeping sickness (human African trypanosomiasis). Tropical Medicine and International Health 12, 1422-1432. 10. Blum J.A., Neumayr A.L., Hatz F. (2011). Human African trypanosomiasis in endemic populations and travellers. European Journal of Clinical Microbiology & Infectious Diseases 31, 905-913. 11. Blum J., Schmid C., Burri C. (2005). Clinical aspects of 2541 patients with second stage human African trypanosomiasis. Acta Tropica 97, 55-64. 12. Brun R., Blum J. (2012). Human African trypanosomiasis. Infectious Disease Clinics of North America 26, 261-273. 13. Brun R., Blum J., Chappuis F., Burri C. (2009). Human African trypanosomiasis. The Lancet 375, 148-159. 14. Buguet A., Bisser S., Josenando T., Chapotot F., Cespuglio R. (2004). Sleep structure: A new diagnostic tool for stage determination in sleeping sickness. Acta Tropica 93, 107-117.

15. Challier A., Gouteux J.P. (1980). Ecology and epidemiological importance of Glossina palpalis in the Ivory Coast forest zone. Insect Science and its Application 1, 77-83. 16. Checchi F., Filipe J.A.N., Barrett M.P., Chandramohan D. (2008). The natural progression of gambiense sleeping sickness: What is the evidence? PLoS Neglected Tropical Diseases 2, e303. 17. Checchi F., Cox A.P., Chappuis F., Priotto F., Chandramohan D., Haydon D.T. (2012). Prevalence and under-detection of gambiense human African trypanosomiasis during mass screening sessions in Uganda and Sudan. Parasites & Vectors 5, 1-13. 18. Clarke J.E. (1964). Game elimination as means of tsetse control with special reference to host preferences. The Puku 2, 67-75. 19. Clarkson M.J. (1976). Trypanosomes. Veterinary Parasitology 2, 9-29. 20. Clausen P.H., Adeyemi I., Bauer B., Breloeer M., Salchow F., Staak C. (1998). Host preferences of tsetse (Diptera: Glossinidae) based on bloodmeal identifications. Medical and Veterinary Entomology 12, 169-180. 21. Clerinx J., Vlieghe E., Asselman V., Van de Casteele S., Maes M.B., Lejon V. (2012). Human African trypanosomiasis in a Belgian traveller returning from the Masai Mara area, Kenya, February 2012. Euro Surveillance 17, 1-4. 22. Cordon-Obras C., Berzosa P., Ndong-Mabale N., Bobuakasi L., Buatiche J.N., Ndongo- Asumu P., Benito A., Cano J. (2009). Trypanosoma brucei gambiense in domestic livestock of Kogo and Mbini foci (Equatorial Guinea). Tropical Medicine and International Health 14, 535- 541. 23. Coustou V., Plazolles N., Guegan F., Baltz T. (2012). Sialidases play a key role in infection and anaemia in Trypanosoma congolense animal trypanosomiasis. Cellular Microbiology 14, 431-445. 24. Eshetu E., Begejo B. (2015). The current situation and diagnostic approach of nagana in Africa: A review. Journal of Natural Sciences Research 5, 117-124. 25. Fèvre E.M., Coleman P.G., Odiit M., Magona J.W., Welburn S.C., Woolhouse M.E.J. (2001). The origins of a new Trypanosoma brucei rhodesiense sleeping sickness outbreak in eastern Uganda. The Lancet 358, 625-628. 26. Franco J.R., Simarro P.P., Diarra A., Jannin J.G. (2014). Epidemiology of human African trypanosomiasis. Clinical Epidemiology 6, 257-275. 27. Geerts S., Holmes P.H., Eisler M.C., Diall O. (2001). African bovine trypanosomiasis: The problem of drug resistance. Trends in Parasitology 17, 25-28. 28. Geigy R., Kauffmann M. (1973). Sleeping sickness survey in the Serengeti Area (Tanzania) 1971. I. Examination of large mammals for trypanosomes. Acta Tropica 30, 12-23. 29. Godrey D., Killick-Kendrick R. (1967). Cyclically transmitted infections of Trypanosoma brucei, T. rhodesiense and T. gambiense in chimpanzees. Transactions of the Royal Society of Tropical Medicine and Hygiene 61, 781-791.

30. Grisard E., Steindel M., Guarneri A.A., Eger-Mangrich I., Campbell D.A., Romanha A.J. (1999). Characterization of Trypanosoma rangeli strains isolated in Central and South America: An overview. Memórias do Instituto Oswaldo Cruz 94, 203-209. 31. Heisch R.B., McMahon J.P., Manson-Bahr P.E.C. (1958). The isolation of Trypanosoma rhodesiense from a bushbuck. British Medical Journal 2, 1203-1204. 32. Herder S., Simo G., Nkinin S., Njiokou F. (2002). Identification of trypanosomes in wild animals from southern Cameroon using the polymerase chain reaction (PCR). Parasite 9, 345-349. 33. Hide G., Tait A., Maudlin I., Welburn S.C. (1996). The origins, dynamics and generation of Trypanosoma brucei rhodesiense epidemics in East Africa. Parasitology Today 12, 50-55. 34. Itard J. (1989). African animal trypanosomoses. In: Shah-Fischer M. and Ralph Say R. (Editors) Manual of tropical veterinary parasitology, CAB international, Wallingford, p. 177- 290. 35. Jahnke H.E., Tacher G., Keil P., and Rojat D. (1988). Livestock production in tropical Africa, with special reference to the tsetse-affected zone. Livestock production in tsetse affected areas of Africa: Proceedings of a meeting in Nairobi, Kenya, 3-21. 36. Jelinek T., Bisoffi Z., Bonazzi L., Van Thiel P., Bronner U., De Frey A., Gundersen S.G., McWhinney P., Ripamonti D. (2002). Cluster of African trypanosomiasis in travelers to Tanzanian national parks. Emerging Infectious Diseases 8, 634-635 37. Jirků M., Votýpa J., Petrželková K.J., Jirků-Pomajbíková K., Kriegová E., Vodička, Lankester F., Leendertz S.A.J., Wittig R.M., Boesch C., Modrý D., Ayala F.J., Leendertz F.H., Lukeš J. (2015). Wild chimpanzees are infected by Trypanosoma brucei. International Journal for Parasitology: Parasites and Wildlife 4, 277-282. 38. Kaare M.T., Picozzi K., Mlengeya T., Fèvre E.M., Mellau L.S., Mtambo M.M., Cleaveland S., Welburn S.C. (2006). Sleeping sickness – A re-emerging disease in the Serengeti? Travel Medicine and Infectious Disease 5, 117-124. 39. Kennedy P.G.E. (2006). Human African trypanosomiasis – Neurological aspects. Journal of Neurology 253, 411-416. 40. Kennedy P.G.E. (2013). Clinical features, diagnosis and treatment of human African trypanosomiasis (sleeping sickness). The Lancet Neurology 12, 186-194. 41. Koffi M., Solano P., Denizot M., Courtin D., Garcia A., Lejon V., Büscher P., Cuny G., Jamonneau V. (2006). Aparasitemic serological suspects in Trypanosoma brucei gambiense human African trypanosomiasis: A potential human reservoir of parasites? Acta Tropica 98, 183-188. 42. Krinsky W.L. (2002). Tsetse Flies (Glossinidae). In: Mullen G. and Durden L. (Editors) Medical and Veterinary Entomology, Academic Press, San Diego, p. 303-316. 43. Kristjanson P.M., Swallow B.M., Rowlands G.J., Kruska R.L., De Leeuw P.N. (1999). Measuring the costs of African animal trypanosomosis, the potential benefits of control and returns to research. Agricultural systems 59, 79-98.

44. Losos G.J., Ikede B.O. (1972). Review of pathology of diseases in domestic and laboratory animals caused by Trypanosoma congolense, T. vivax, T. brucei, T. rhodesiense and T. gambiense. Veterinary Pathology Supplementum 9, 1-71. 45. Maes L. (2014). Cursus Algemene Parasitologie, Faculteit Farmaceutische, Biomedische en Diergeneeskundige Wetenschappen, Antwerpen. 46. Malvy D., Chappuis F. (2011). Sleeping sickness. Clinical Microbiology and Infection 17, 986- 995. 47. Molyneux D.H., Ashford R.W. (1989). African human trypanosomiases; African animal trypanosomiasis. In: Molyneux D.H. and Ashford R.W. (Editors) The biology of Trypanosoma and Leishmania, parasites of man and domestic animals, Taylor and Francis, London, p. 97- 151. 48. Mulla A.F., Rickman L.R. (1988). How do African game animals control trypanosome infections? Parasitology Today 4, 352-354. 49. Murray M., Clifford D.J., Gettinby G., Snow W.F., McIntyre W.I. (1981). Susceptibility to African trypanosomiasis of N’Dama and Zebu cattle in an area of Glossina morsitans submorsitans challenge. Veterinary Record 109, 503-510. 50. Naessens J. (2005). Bovine trypanotolerance: A natural ability to prevent severe anaemia and haemophagocytic syndrome? International Journal for Parasitology 36, 521-528. 51. Namangala B., Odongo S., (2014). Animal African trypanosomosis in Sub-Saharan Africa and beyond African borders. In: Magez S. and Radwanska M. (Editors). Trypanosomes and trypanosomiasis, Springer-Verlag, Vienna, p. 239-260. 52. Njiokou F., Laveissière C., Simo G., Nkinin S., Grébaut P., Cuny G., Herder S. (2005). Wild animal as a probable animal reservoir for Trypanosoma brucei gambiense in Cameroon. Infection, Genetics and Evolution 6, 147-153. 53. Njiokou F., Nimpaye H., Simo G., Njitchouang G.R., Asonganyi T., Cuny G., Herder S. (2010). Domestic animals as potential reservoir hosts of Trypanosoma brucei gambiense in sleeping sickness foci in Cameroon. Parasite 17, 61-66. 54. Odiit M., Kansiime F., Enyaru J.C. (1997). Duration of symptoms and case fatality of sleeping sickness caused by Trypanosoma brucei rhodesiense in Tororo, Uganda. East African Medical Journal 74, 792-795. 55. Onyango R.J., Van Hoeve K., De Raadt P. (1966). The epidemiology of Trypanosoma rhodesiense sleeping sickness in Alego location, Central Nyanza, Kenya. I. Evidence that cattle may act as reservoir hosts of trypanosomes infective to man. Transactions of the Royal Society of Tropical Medicine and Hygiene 60, 175-182. 56. Picozzi K., Fèvre E.M., Odiit M., Carrington M., Eisler M.C., Maudlin I., Welburn S.C. (2005). Sleeping sickness in Uganda: A thin line between two fatal diseases. The BMJ 331, 1238- 1242. 57. Radwanska M., Chamekh M., Vanhamme L., Claes F., Magez S., Magnus E., De Baetselier P., Büscher P., Pays E. (2002). The serum resistance-associated gene as a diagnostic tool for the

detection of Trypanosoma brucei rhodesiense. American Journal of Tropical Medicine and Hygiene 67, 684-690. 58. Roditi I., Lehane M.J., (2008). Interactions between trypanosomes and tsetse flies. Current Opinion in Microbiology 11, 345-351. 59. Rotureau B., Van Den Abbeele J. (2013). Through the dark continent: African trypanosome development in the tsetse fly. Frontiers in Cellular and Infection Microbiology 3, 1–7. 60. Saini R.K., Hassanali A. (2007). A 4-alkyl-substituted analogue of guaiacol shows greater repellency to savannah tsetse (Glossina spp.). Journal of Chemical Ecology 33, 985-995. 61. Sbicego S., Vassella E., Kurath U., Blum B., Roditi I. (1999). The use of transgenic Trypanosoma brucei to identify compounds inducing the differentiation of bloodstream forms to procyclic forms. Molecular and Biochemical Parasitology 104, 311-322 62. Simarro P.P., Diarra A., Postigo J.A.R., Franco J.R., Jannin J.G. (2011). The human African trypanosomiasis control and surveillance programme of the World Health Organization 2000- 2009: The way forward. PLoS Neglected Tropical Diseases 5, e1007. 63. Simo G., Asonganyi T., Nkinin S.W., Njiokou F., Herder S. (2006). High prevalence of Trypanosoma brucei gambiense group 1 in pigs from the Fontem sleeping sickness focus in Cameroon. Veterinary Parasitology 139, 57-66. 64. Simukoko H., Marcotty T., Phiri I., Geysen D., Vercruysse J., Van den Bossche P. (2007). The comparative role of cattle, goats and pigs in the epidemiology of livestock trypanosomiasis on the plateau of eastern Zambia. Veterinary Parasitology 147, 231-238. 65. Swallow B.M. (1999). Impacts of African animal trypanosomosis on migration, livestock and crop production. International Scientific Council of Trypanosomiasis Research and Control, International Livestock Research Institute, Nairobi, Kenya. 66. Tirados I., Esterhuizen J., Rayaisse J.B., Diarrassouba A, Kaba D., Mpiana S., Vale G.A., Solano P., Lehane M.J., Torr S.J. (2011). How do tsetse recognise their hosts? The role of shape in the responses of tsetse (Glossina fuscipes and G. palpalis) to artificial hosts. PLoS Neglected Tropical Diseases 5, e1226. 67. Tongue L.K., Mavoungou J.F., Kamkumo R.G., Kaba D., Hendji G.C.F., Pamba R., M’eyi P.M., Mbatchi B., Louis F. (2012). Human African trypanosomiasis in suburban and urban areas: A potential challenge in the fight against the disease. Journal of Clinical & Experimental Pathology S3, 1-5. 68. Uilenberg G. (1998). A field guide for the diagnosis, treatment and prevention of African animal trypanosomosis. Food and Agriculture Organization of the United Nations. 69. Urech K., Neumayr A., Blum J. (2011). Sleeping sickness in travelers – Do they really sleep? PLos Neglected Tropical Diseases 5, e1358. 70. Uzureau P., Uzurea S., Lecordier L., Fontaine F., Tebabi P., Homblé F., Grélard A., Zhendre V., Nolan D.P., Lins L., Crowet J.M., Pays A., Felu C., Poelvoorde P., Vanhollebeke B., Moestrup S.K., Lyngsø J., Pedersen J.S., Mottram J.C., Dufourc E.J., Pérez-Morga D., Pays E. (2013). Mechanism of Trypanosoma brucei gambiense resistance to human serum. Nature 501, 430-434.

71. Van den Bossche P., Rowlands G.J. (2000). The relationship between the parasitological prevalence of trypanosomal infections in cattle and herd average packed cell volume. Acta Tropica 78, 163-170. 72. Van Hoeve K., Onyango R.J., Harley J.M.B., De Raadt P. (1967) The epidemiology of Trypanosoma rhodesiense sleeping sickness in Alego location, Central Nyanza, Kenya. II. The cyclical transmission of Trypanosoma rhodesiense isolated from cattle to a man, a cow and to sheep. Transactions of the Royal Society of Tropical Medicine and Hygiene 61, 684-687. 73. Vickerman K., Tetley L., Hendry K.A.K., Turner C.M.R. (1988). Biology of African trypanosomes in the tsetse fly. Biology of the Cell 64, 109-119. 74. Vickerman K. (1985). Developmental cycles and biology of pathogenic trypanosomes. British Medical Bulletin 41, 105-114. 75. Weitz B. (1963). The feeding habits of Glossina. Bulletin of the World Health Organization 28, 711-729. 76. Welburn S.C., Picozzi K., Fèvre E.M., Coleman P.G., Odiit M., Carrington M., Maudlin I. (2001). Identification of human-infective trypanosomes in animal reservoir of sleeping sickness in Uganda by means of serum-resistance-associated (SRA) gene. The Lancet 358, 2017- 2019. 77. Wilson S.G., Morris K.R.S., Lewis I.J., Krog E. (1963). The effects of trypanosomiasis on rural economy: With special reference to Sudan, Bechuanaland and West Africa. Bulletin of the World Health Organization 28, 595-613. 78. World Health Organization. (2013). Control and surveillance of Human African trypanosomiasis. WHO Technical Report Series 984.

ANNEXES Annex 1: Species and subspecies of Glossina. Source: WHO (2013).

* Major vectors of sleeping sickness