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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
by
Kim VAN DE WIEL
Promoter: Prof. Dr. Pierre Dorny
Co-promoter: Prof. Dr. Louis Maes
Literature Review
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
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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
Co-promoter: Prof. Dr. Louis Maes
Literature Review
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
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
1
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
2
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 subSaharan 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.
3
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
4
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 subdermally, 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
5
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
6
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).
7
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
8
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
A: 38.8%
A: 18.2%
A: 42.7%
B: 18.2%
B: 8.9%
B: 2.0%
A: 5.5%
A: 3.2%
A: 1.9%
B: 43,8%
B: 15.3%
B: 0.6%
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%
Primates
Suids
Ruminants
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
A*: 10.4%
A: 1.9%
A: 2.7%
A: 4.9%
B**: 0.7%
B: 2.5%
B: 2.3%
B: 5.2%
A*: 36.4%
A: 4.3%
A: 29.9%
A: 57.7%
B**: 57.1%
B: 10.2%
B: 36.2%
B: 89.7%
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%
Primates
Suids
Ruminants
*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
9
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
A: 0%
A: 0.7%
A: 0.7%
A: 0.4%
B: 0%
B: 1.4%
B: 0%
B: 0.25%
A: 13.7%
A: 64.4%
A: 39.4%
A: 1.0%
B: 7.6%
B: 70.7%
B: 8.6%
B: 60.6%
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%
Primates
Suids
Ruminants
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).
10
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).
11
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 km 2, are at risk of contracting sleeping
sickness caused by T. b. gambiense. Approximately 12.3 million people in a region of 0.171 million km 2
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 meningoencephalitic 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.,
12
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 nonendemic 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).
13
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
14
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).
15
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 manfly-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
16
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.
17
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.
18
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
19
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
20
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.,
21
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.
22
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.
23
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ANNEXES
Annex 1: Species and subspecies of Glossina. Source: WHO (2013).
* Major vectors of sleeping sickness
30