GHENT UNIVERSITY

FACULTY OF VETERINARY MEDICINE

Academic year 2015-2016

THE IMPORTANCE OF WILDLIFE AS A RESERVOIR

FOR HUMAN AND ANIMAL AFRICAN TRYPANOSOMIASIS

by

Kim VAN DE WIEL

Promoter: Prof. Dr. Pierre Dorny Literature Review

Co-promoter: Prof. Dr. Louis Maes as part of the Master’s Dissertation

© 2016 Kim van de Wiel

Disclaimer:

Universiteit Gent, its employees and/or students, give no warranty that the information provided in this thesis is accurate or exhaustive, nor that the content of this thesis will not constitute or result in any infringement of third-party rights.

Universiteit Gent, its employees and/or students do not accept any liability or responsibility for any use which may be made of the

content or information given in the thesis, nor for any reliance which may be placed on any advice or information provided in this thesis.

GHENT UNIVERSITY

FACULTY OF VETERINARY MEDICINE

Academic year 2015-2016

THE IMPORTANCE OF WILDLIFE AS A RESERVOIR

FOR HUMAN AND ANIMAL AFRICAN TRYPANOSOMIASIS

by

Kim VAN DE WIEL

Promoter: Prof. Dr. Pierre Dorny Literature Review

Co-promoter: Prof. Dr. Louis Maes as part of the Master’s Dissertation

© 2016 Kim van de Wiel

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

1

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

2

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

3

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.

4

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

5

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

6

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

7

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).

8

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

9

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%

B: 18.2%

A: 18.2%

B: 8.9%

A: 42.7%

B: 2.0%

Suids A: 5.5%

B: 43,8%

A: 3.2%

B: 15.3%

A: 1.9%

B: 0.6%

Ruminants A: 22,0%

B: 17.8%

A: 37.8%

B: 22.9%

A: 30.4%

B: 33.6%

Other

mammals

A: 4.1%

B: 7.0%

A: 5.1%

B: 6.5%

A: 16.0%

B: 49.7%

Reptiles A: 27.7%

B: 10.5%

A: 34.4%

B: 42.1%

A: 8.3%

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%

B**: 0.7%

A: 1.9%

B: 2.5%

A: 2.7%

B: 2.3%

A: 4.9%

B: 5.2%

Suids A*: 36.4%

B**: 57.1%

A: 4.3%

B: 10.2%

A: 29.9%

B: 36.2%

A: 57.7%

B: 89.7%

Ruminants A*: 45.2%

B**: 21%

A: 91.5%

B: 72.8%

A: 63.5%

B: 52.2%

A: 35.6%

B: 3.4%

Other

mammals

A*: 7.0%

B**: 20.7%

A: 2.2%

B: 3.9%

A: 3.5%

B: 8.2%

A: 1.8%

B: 0%

Reptiles A*: 0.3%

B**: 0.2%

A: 0%

B: 9.5%

A: 0.2%

B: 0.6%

A: 0%

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

10

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%

B: 0%

A: 0.7%

B: 1.4%

A: 0.7%

B: 0%

A: 0.4%

B: 0.25%

Suids A: 13.7%

B: 7.6%

A: 64.4%

B: 70.7%

A: 39.4%

B: 8.6%

A: 1.0%

B: 60.6%

Ruminants A: 73.6%

B: 84.0%

A: 19.9%

B: 21.2%

A: 23.4%

B: 6.4%

A: 17.9%

B: 21.5%

Other

mammals

A: 12.9%

B: 4.2%

A: 15.0%

B: 6.6%

A: 36.3%

B: 85.0%

A: 73.4%

B: 15.6%

Reptiles A: 0%

B: 3.4%

A: 0%

B: 0%

A: 0%

B: 0%

A: 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).

11

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).

12

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.,

13

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).

14

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

15

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).

16

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

17

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.

18

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.

19

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

20

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

21

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.,

22

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.

23

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.

24

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.

25

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.

26

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.

27

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

28

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.

29

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.

30

ANNEXES

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

* Major vectors of sleeping sickness