Nutritional stress affects the tsetse fly's immune gene expression

i

Effect of nutritional stress on the tsetse fly’s vector competence and its

implications on trypanosome transmission in the field

Komlan AKODA

Dissertation submitted in fulfilment of the requirements for the degree of Doctor

(PhD) in Veterinary Sciences

2009

Promoters:

Prof. Dr. P. Van den Bossche

Prof. Dr. Pierre Dorny

Co-Promoters:

Dr. Jan Van Den Abbeele

Dr. Issa Sidibé

Laboratory of Parasitology

Department of Virology, Parasitology and Immunology

Faculty of Veterinary Medicine, Ghent University

Salisburylaan 133, B-9820 Merelbeke

i

‘‘Gloire à toi Dieu le Père, le Fils et le Saint Esprit pour m’avoir donné

la vie et la chance d’accomplir ce travail’’

I dedicate this thesis to my mother, my uncle and my late father

ii

Acknowledgments

First and foremost, the most important acknowledgment goes to my ITM promoters Dr. Jan

Van Den Abbeele and Prof. Dr. Peter Van den Bossche for initiating this research project and

for their invaluable and constructive guidance and tireless support during this thesis

I would also like to express my profound gratitude to Prof. Dr. Pierre Dorny for accepting me

as his student at Ghent University

My sincere gratitude also goes to the Belgiun government through the DGDC (Directory

General of Development Cooperation) and the Institute of Tropical Medicine (ITM) for the

financial support of this thesis

Many thanks to my local promotor Dr. Issa Sidibé, the Scientific Director of CIRDES (Bobo-

Dioulasso/ Burkina Faso), for facilities he provided to me during my field work in Burkina

Faso

I would also like to extend my acknowledgment to Prof. Louis Joseph Pangui, the Director of

EISMV (Dakar, Senegal) and Dr. Assiongbon Teko-Agbo for allowing me to accomplish this

thesis

Special thanks go to Dr. Tanguy Marcotty for his tireless efforts and patience in helping me

with the statistical analysis of my experimental data

I want to say “Bedankt” to Karin De Ridder and Jos Van Hees of the Entomology Section in

the Parasitology Department for their technical assistance

My sincere thanks go to all the members of staff of the Animal Health Department of the

Institute of Tropical Medicine, Antwerp. More specifically to Prof. Dr. Stanny Geerts, Dr.

Vincent Delespaux, Dr. Redgi De Deken, Mrs Danielle De Bois and all the technical staff

I thank Dr. Sophie Ravel at IRD-CIRAD (Montpellier/France) and her technicians for their

assistance during my experiments in their laboratory

I acknowledge all people who assist me in different ways during this work, especially Dr.

Zakaria Bengaly and the technicians of the laboratory of Parasitology in CIRDES (Bobo-

Dioulasso/Burkina Faso), the staff of Rekomitjie Research Station in Zimbabwe

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Thanks are due to my colleagues and/or friends Alain Nahum, Linda De Vooght, Nynke

Deckers, Simbarashe Chitanga, Emmanel Abatih, Emmanuel Assana, Evelyne Houdjè,

Moussa Sanogo, Alexis Koffi Doua, Jérome Nkuna and Eugène Ubertalli

A big thank to my lovely Bénédicte Amsatou Darga, whose understanding, constant

encouragement and support really kept me going. Could this thesis be a reward of your

patience and support

Lastly but not the least I thank my family for their support

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List of Abbreviations AAT: African Animal Trypanosomiasis

AMP: Antimicrobial peptide

cDNA: Complementary Deoxyribonucleic acid

dNTP: Deoxynucleoside triphosphates

E. coli: Escherichia coli

FAO: Food and Agriculture Organization

HAT: Human African Trypanosomiasis

H2O2: Hydrogen Peroxide

IVC: Intrinsic Vectorial Capacity

mL: Milliliter

µL: Microliter

mRNA: messenger Ribonucleic acid

NO Nitric Oxide

PBS: Phosphate Buffered Saline

PCR: Polymerase Chain Reaction

ROS: Reactive Oxygen Species

RTQ-PCR: Real time quantitative PCR

STATAcorp: Stata Corporation statistical software

WHO: World Health Organization

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List of Figures Number Title Page

Figure 1 Classification of human and animal pathogenic tsetse-transmitted trypanosome species………………………………………………...

3

Figure 2 Distribution and number of tsetse fly species belonging to the fusca, morsitans and palpalis groups in Africa…………………………….

5

Figure 3 Life cycle of Trypanosoma brucei………………………………….. 7

Figure 1.1 Diagram illustrating different key stages of Trypanosoma brucei development in the tsetse fly and the mammal host………………...

12

Figure 1.2 Tsetse fly–related and exogenous factors interfering with the tsetse-trypanosome interaction……………………………………………..

13

Figure 1.3 The molecular components involved in the two major signalling pathways in the Drosophila immune response……………………...

17

Figure 4.1 Normalized expression levels of attacin, defensin and cecropin in male teneral G. m. morsitans emerging from non-starved or starved adult female flies and their 95% confidence intervals (n = 24 and 20 abdomens respectively in non-starved and starved flies group)…….

54

Figure 5.1 Fat content (± 95% confidence intervals; n=12 per group) of male teneral and 20-days-old G. m. morsitans after a period of starvation.

67

Figure 5.2 Normalized expression levels (± 95% confidence intervals; n=12 per group) of attacin (dark grey bar), defensin (white bar) and cecropin (light grey bar) in male teneral and 20-days-old G. m.

morsitans after starvation. Expression levels (mRNA) of attacin, defensin and cecropin were measured in non-starved (TF0) and starved (TF4) teneral (A) and non-starved (AD2) and starved (AD7) 20-days-old flies (B) using quantitative real-time PCR……..

68

Figure 5.3 Normalized expression levels (± 95% confidence intervals; n=4 per group) of attacin (dark grey bar), defensin (white bar) and cecropin (light grey bar) in non- starved (TF0) and starved (TF4) teneral (A) and in non-starved (AD2) and starved (AD7) 20-days-old (B) male G. m. morsitans after bacterial (E. coli) challenge…………………..

69

Figure 5.4 Normalized gene expression levels (± 95% confidence intervals; n=5 per group) of attacin (dark grey bar), defensin (white bar) and cecropin (light grey bar) in male non-starved (TF0) and starved (TF4) teneral (A) or in male non-starved (AD2) and starved (AD7) 20-days-old flies (B) G. m. morsitans on day 1 (d1) or day 5 (d5) after the uptake of a blood meal containing T. congolense (Tc) blood stream forms…………………………………………………..

70

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Figure 5.5

Normalized gene expression levels (± 95% confidence intervals; n=5 per group) of attacin (dark grey bar), defensin (white bar) and cecropin (light grey bar) in male non-starved (TF0) and starved (TF4) teneral (A) or in male non-starved (AD2) and starved (AD7) 20-day-old flies (B) G.m. morsitans on day 1 (d1) or day 5 (d5) after the uptake of a blood meal containing T. brucei brucei (Tb) bloodstream forms…………………………………………………...

71

Figure 6.1 A map of Zimbabwe showing the Zambezi Valley and the sampling area ………………………………………………………………….

79

Figure 6.2 Fat body content (± 95% confidence intervals) of male non-teneral G. m. morsitans (n= 100) and G. pallidipes (n=50) collected during a rainy season and hot dry season in the field……………………….

81

Figure 6.3 Normalized expression levels (± 95% confidence intervals; n=30 per group) of attacin, defensin and cecropin in male non-teneral G.

m. morsitans (A) and G. pallidipes (B) collected during the rainy season and the hot dry season in the Zambezi Valley of Zimbabwe..

82

Figure 7.1 Average proportion of mature infection of male G. m. morsitans infected on different days post-infection on mice infected with T.

congolense IL1180…………………………………………………..

92

Figure 8.1 Diagram of parameters affecting fat depletion and immunosuppression in tsetse flies…………………………………...

99

Figure 8.2 Map of the satellite measured average maximum temperature (A) and location of tsetse belts, populated places and T. b. rhodesiense historical foci (B)……………………………………………………

101

Figure 8.3 Monthly average fat body levels and T. congolense infection rates in G. m. morsitans in the Eastern Zambia …………………………..

102

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List of Tables Number Title Page

Table 1 Susceptibility of livestock species to the pathogenic trypanosome species……………………………………………………………………...

2

Table 2 Characterization of trypanosomes according to their site of development in Glossina spp…………………………………………………………......

2

Table 2.1 Proportion (+95% confidence interval) of starved and non-starved teneral and 20-day-old G. m. morsitans male flies that developed a trypanosome infection with T. b. rhodesiense in the midgut (procyclic stage) and salivary glands (mature, metacyclic stage), 30 days after the infected blood meal………………………………………………………………….

37

Table 2.2 Significance (P value) of the difference between the proportion of male G.

m. morsitans with a procyclic or metacyclic infection and IVC compared to non-starved flies (Group 1 and Group 4 served as reference for teneral and older flies, respectively)……………………………………………….

37

Table 3.1 Proportion (+95% confidence interval) of G. m. morsitans male flies that developed a trypanosome infection with T.brucei brucei AnTAR 1 in the midgut (procyclic stage) and salivary glands (mature, metacyclic stage), 30 days after the infected blood meal………………………………………

46

Table 4.1 Proportion of male teneral G. m. morsitans emerging from starved or non-starved females and infected with T. congolense IL1180………………….

55

Table 4.2 Proportion of male teneral G. m. morsitans emerging from starved or non-starved females and infected with T. b. brucei AnTAR 1…………………

56

Table 6.1 Number and proportion (%) of non-teneral (male and females) tsetse G.

m. morsitans and G. pallidipes infected with trypanosomes in the rainy season and the hot dry season………………………………………………

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Table 7.1 Immature and mature infection rates of male G. m. morsitans given a single infective bloodmeal on various days of the development of T.

congolense IL1180 in mice…………………………………………….......

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Table 7.2 Significance (P value) of the difference between the proportion of male G.

m. morsitans with a mature or immature infection and infected on days 5, 6, 7 or 10 post-infection compared to flies infected on day 4 post-infection (reference)……………………………………………………………….....

92

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Table of Contents Acknowledgments ................................................................................................................................................. ii

List of abbreviations ............................................................................................................................................ iv

List of Figures........................................................................................................................................................ v

List of Tables ....................................................................................................................................................... vii

Table of Contents ............................................................................................................................................... viii

General introduction............................................................................................................................................. 1

1. Introduction....................................................................................................................................................... 1

2. The parasite: Trypanosoma spp ........................................................................................................................ 1

3. The vector: Glossina spp................................................................................................................................... 3

3.1. Morphological features and classification................................................................................................... 3 3.2. Life cycle..................................................................................................................................................... 4 3.3. Distribution and habitat preference ............................................................................................................. 4

4. Life cycle of trypanosomes ............................................................................................................................... 6

5. Impact of trypanosomiasis on animal and human health.............................................................................. 7

6. Trypanosomiasis control .................................................................................................................................. 8

6.1. Control of the parasite in the host................................................................................................................ 8 6.2. Control of the vector.................................................................................................................................... 8

Chapter 1 ............................................................................................................................................................. 10

Factors affecting the tsetse’s susceptibility to trypanosome infection: A literature review.......................... 10

1.1. Establishment and migration/maturation of trypanosome infection....................................................... 11

1.2. Endogenous factors affecting tsetse-trypanosome interaction ................................................................. 14

1.2.1. Molecular base interfering with trypanosome infection in tsetse ........................................................... 14 1.2.1.1. Role of lectins ................................................................................................................................. 14 1.2.1.2 Role of tsetse EP- proteins............................................................................................................... 15 1.2.1.3. Role of tsetse flies midgut proteases ............................................................................................... 15 1.2.1.4. Role of tsetse immune system.......................................................................................................... 15

1.2.2. Role of endosymbionts in the development of a trypanosome infection................................................ 19

1.3. Exogenous factors affecting tsetse-trypanosome interaction.................................................................... 19

1.3.1. Temperature ........................................................................................................................................... 19 1.3.2. Role of host blood .................................................................................................................................. 20 1.3.3. Role of tsetse fly starvation .................................................................................................................... 20 1.3.4. Role of trypanosome genotype............................................................................................................... 21

1.4. Conclusion .................................................................................................................................................... 21

1.5. References..................................................................................................................................................... 23

Objectives of the thesis ....................................................................................................................................... 30

Chapter 2 ............................................................................................................................................................. 32

Effect of starvation on the tsetse fly’s susceptibility to human pathogenic trypanosome species (T. b.

rhodesiense and T. b. gambiense) ....................................................................................................................... 32

2.1. Introduction.................................................................................................................................................. 33

2.2. Material and methods .................................................................................................................................. 33

2.2.1. Trypanosome isolates ............................................................................................................................. 33 2.2.2. Tsetse flies and experimental groups ..................................................................................................... 34 2.2.3. Tsetse flies infective blood meal and maintenance ................................................................................ 35 2.2.4. Tsetse flies dissection and infection rates............................................................................................... 35 2.2.5. Statistical analysis .................................................................................................................................. 35

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2.3. Results ........................................................................................................................................................... 36

2.3.1. Infections with T. b. gambiense ............................................................................................................. 36 2.3.2. Infections with T. b. rhodesiense............................................................................................................ 36

2.4. Discussion...................................................................................................................................................... 37

2.5. References..................................................................................................................................................... 40

Chapter 3 ............................................................................................................................................................. 42

Maturation of a Trypanosoma brucei infection to the infectious metacyclic stage is enhanced in

nutritionally stressed tsetse flies ........................................................................................................................ 42

3.1. Introduction.................................................................................................................................................. 43

3.2. Materials and Methods................................................................................................................................ 43

3.2.1. Tsetse flies and trypanosome strain........................................................................................................ 43 3.2.2. Experimental design ............................................................................................................................... 44

3.3. Results and Discussion................................................................................................................................. 45

3.4. References..................................................................................................................................................... 48

Chapter 4 ............................................................................................................................................................. 50

Nutritional stress of adult female tsetse flies (Diptera: Glossinidae) affects the susceptibility of their

offspring to trypanosomal infections ................................................................................................................. 50

4.1. Introduction.................................................................................................................................................. 51

4.2. Materials and methods ................................................................................................................................ 52

4.2.1. Tsetse flies.............................................................................................................................................. 52 4.2.2. Trypanosome infection of tsetse flies..................................................................................................... 52 4.2.3. Tsetse infection rate................................................................................................................................ 52 4.2.4. Determination of the fat content in the offspring ................................................................................... 53 4.2.5. Immune peptide gene expression levels in the freshly emerged flies..................................................... 53 4.2.6. Statistical data analysis........................................................................................................................... 53

4.3. Results ........................................................................................................................................................... 53

4.3.1. Nutritional stress and expression of immune peptide genes of the emerging flies................................. 54 4.3.2. Trypanosome infection rates .................................................................................................................. 54

4.4. Discussion...................................................................................................................................................... 56

4.5. References..................................................................................................................................................... 59

Chapter 5 ............................................................................................................................................................. 62

Nutritional stress affects the tsetse fly’s immune gene expression.................................................................. 62

5.1. Introduction.................................................................................................................................................. 63


5.2.1. Tsetse flies.............................................................................................................................................. 64 5.2.2. Nutritional stress by starvation............................................................................................................... 64 5.2.3. Fat content determination....................................................................................................................... 65 5.2.4. Bacterial and trypanosomal challenge of tsetse flies .............................................................................. 65 5.2.5. RNA extraction and quantification......................................................................................................... 65 5.2.6. First- strand cDNA synthesis ................................................................................................................. 66 5.2.7. Quantitative Real - time PCR................................................................................................................. 66 5.2.8. Statistical data analysis........................................................................................................................... 66

5.3. Results ........................................................................................................................................................... 67

5.3.1. Fat content and nutritional stress ............................................................................................................ 67 5.3.2. Immune gene expression level and nutritional stress ............................................................................. 68 5.3.3. Immune gene response to bacterial/trypanosomal challenge and nutritional stress ............................... 69

5.4. Discussion...................................................................................................................................................... 72

5.5. References..................................................................................................................................................... 75

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Chapter 6 ............................................................................................................................................................. 77

Investigation of the effect of seasonal climatic change on nutritional and immune status and trypanosome

infection in natural field-caught tsetse flies ...................................................................................................... 77

6.1. Introduction.................................................................................................................................................. 78


6.2.1. Study site ................................................................................................................................................ 79 6.2.2. Trapping of tsetse and sample collection ............................................................................................... 80 6.2.3. Fly dissection.......................................................................................................................................... 80 6.2.4. Trypanosome species identification ....................................................................................................... 80 6.2.5. Fat body content and immune peptide expression analysis .................................................................... 80 6.2.6. Statistical data analysis........................................................................................................................... 81

6.3. Results ........................................................................................................................................................... 81

6.3.1. Fat body content and immune peptide expression levels ....................................................................... 81 6.3.2. Tsetse infection rates and trypanosome species ..................................................................................... 83

6.4. Discussion...................................................................................................................................................... 83

6.5. Reference ...................................................................................................................................................... 86

Chapter 7 ............................................................................................................................................................. 88

Investigations on the transmissibility of Trypanosoma congolense by the tsetse fly Glossina morsitans

morsitans during its development in a mammalian host.................................................................................. 88

7.1. Introduction.................................................................................................................................................. 89


7.2.1. Tsetse flies.............................................................................................................................................. 89 7.2.2. Trypanosome.......................................................................................................................................... 90 7.2.3. Experimental design ............................................................................................................................... 90

7.3. Results ........................................................................................................................................................... 91

7.4. Discussion...................................................................................................................................................... 93

7.5. References..................................................................................................................................................... 95

Chapter 8 ............................................................................................................................................................. 97

General discussion............................................................................................................................................... 97

8.1. Introduction.................................................................................................................................................. 98

8.2. Tsetse starvation and trypanosome development...................................................................................... 98

8.3. Factors inducing nutritional stress in tsetse flies....................................................................................... 99

8.3.1. Ambient temperature (climate)............................................................................................................. 100 8.3.1.1. Geographical location (latitude) ........................................................................................................ 100 8.3.1.2. The altitude........................................................................................................................................ 100 8.3.1.3. Seasonality ........................................................................................................................................ 102 8.3.1.4. Climate change .................................................................................................................................. 103 8.3.2. Availability of hosts ............................................................................................................................. 103

8.4. References................................................................................................................................................... 104

Summary............................................................................................................................................................ 105

Samenvatting ..................................................................................................................................................... 109

General introduction


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1. Introduction

African trypanosomiasis is one of the most important parasitic diseases caused by several

species of trypanosomes. In 36 countries of sub-Saharan Africa, tsetse-transmitted

trypanosomes affect both humans and animals. In humans, the disease called sleeping

sickness causes a considerable public health burden on rural populations (Barret, 1999). It is

estimated that about 70 million people on the African continent are at risk with about 50 to 70

thousands of people infected annually (WHO, 2006). Animal trypanosomosis (nagana) is an

important constraint to livestock production in sub-Saharan Africa, preventing full use of the

land to feed the rapidly increasing human population. Not controlled, the disease can induce

important losses by limiting crop production due to less efficient nutrient cycling, reduced

access to animal traction, lower income from milk and meat sales and reduced access to liquid

capital (Swallow, 2000). The Food and Agriculture Organisation (FAO) attributes 3 million

cattle deaths to trypanosomiasis and current losses in cattle production alone amounting to

between US$ 1.0 – 1.2 billion annually (DFID, 2001).

2. The parasite: Trypanosoma spp

Trypanosomes are flagellated protozoan parasites belonging to the order Kinetoplastida,

family Trypanosomatidae, genus Trypanosoma (Hoare, 1972). They are extracellular parasites

mostly found in the blood circulation of their mammalian host. However, tissue localisation

has also been observed in some species such as T. brucei spp., T. vivax and T. equiperdum

(Stephen, 1986). Two T. brucei subspecies are pathogenic for humans and differ in virulence

and geographical distribution. Trypanosoma brucei gambiense causes sleeping sickness in

West and Central Africa and has a chronical course of infection while T. b. rhodesiense

causes an acute infection in humans in East and southern Africa. In the latter case, the disease

is zoonotic with both wild and domestic animals being important reservoir of the parasite.

Several other trypanosome species are animal pathogenic. Susceptibility of livestock to a

trypanosome infection varies depending on the trypanosome species, with some livestock

species being more susceptible to one or more trypanosome species than others (Table 1).


2

Table 1: Susceptibility of livestock species to the pathogenic trypanosome species (Adapted

from Soltys, 1963)

Trypanosome species Livestock species T. congolense T. simiae T. vivax T. brucei T. evansi T. equiperdum

T.

suis

Cattle +++ ± ++ + + � � Sheep ++ + ++ +++ ++ � � Goat ++ + ++ +++ ++ � � Pig + +++ � + � � +++ Horse ++ � ++ +++ ++ + � Camel ++ � � ++ ++ � �

+++: very susceptible, ++: susceptible, +: less susceptible, �: no infection

Tsetse-transmitted trypanosomes belong to the section of the Salivaria. Based on several

criteria among which the parasite’s morphology and the site of development in the tsetse fly

vector (Table 2), trypanosomes are classified, within this section, into four different subgenera

and furthermore into different species (Figure 1).

Table 2: Characterization of trypanosomes according to their site of development in Glossina

spp. After Hoare (1970) and Van Den Abbeele et al. (1999).

Subgenus Species Midgut Proboscis salivary glands

Duttonella T.vivax No development Trypomastigotes No development

epimastigotes

Final infective stage: metacyclic trypomastigotes

Nannomonas T.congolense Trypomastigotes Trypomastigotes No development

T.simiae epimastigotes

T.godfreyi

Final stage: metacyclic trypomastigotes

Trypanozoon T.b.brucei Trypomastigotes Trypomastigotes

and epimastigotes Trypomastigotes and epimastigotes

T.b.rhodesiense

Final stage: metacyclic trypomastigotes

T.b. gambiense


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3. The vector: Glossina spp.

3.1. Morphological features and classification

Tsetse flies are obligate blood feeding insects with both male and female taking blood meals

every two to five days contributing hence to trypanosome transmission. They belong to the

order of the Diptera, family Glossinidae, genus Glossina. The genus Glossina is subdivided in

three groups or subgenera, namely the fusca group, the morsitans group and the palpalis

group (Buxton, 1955). Each group is subdivided into species and subspecies. Up to now, there

are 31 identified species/subspecies of tsetse flies.

Figure 1: Classification of human and animal pathogenic tsetse-transmitted trypanosome

species (Masumu Mulumba, 2006)

Su

bsp

ecies/Su

bg

rou

ps

Sp

ecies S

ub

gen

us

Sectio

n

Gen

us

Dutonella Nannomonas Pycnomonas

T. vivax T. congolense

T. simiae

T. godfreyi

T. brucei s.l

T. evansi

T. equiperdum

T. suis

Trypanosoma

Salivaria Stercoraria (spp : T. theileri)

Trypanozoon

T. congolense Savannah T. congolense WARF T. congolense Kilifi T. congolense Tsavo

T. b. brucei

T. b. gambiense

T. b. rhodesiense


4

Morphologically, tsetse flies are rather dull in appearance, varying in colour from a

light yellowish-brown to a dark blackish-brown. The abdomen may be uniformly coloured or

transversed by stripes according to species. Two visually distinctive characteristics that are

apparent to the eye are the forward projecting proboscis and the unusual hatchet shaped cell

formed by the wing venation.

3.2. Life cycle

Tsetse flies reproduce by adenotrophic viviparity i.e. giving birth to live offspring nourished

within the mother by secretion from highly modified accessory glands and born at an

advanced stage of development. The female tsetse fly is inseminated only once during the

course of its reproductive life and produces offspring at regular interval. It produces a single

egg, which hatches into a first-stage larva in the uterus. After a period of development and

moulting, a third-larva is deposited on the ground. The three larval stages develop within the

mother. They are nourished with a milky secretion produced by the female accessory glands.

Females produce one full-grown larva every 9-10 days, which then pupates within 1-2 hours.

The adult fly will emerge after a puparial period that varies according to temperature but is

about 30 days at 24oC (Leak, 1999). The newly emerged fly that has not taken its first blood

meal is called teneral fly.

3.3. Distribution and habitat preference

Tsetse flies are found exclusively in about 10 million km2 of sub-Saharan Africa. This area,

ranging between 14°N and 29°S from Senegal in the West to Southern Somalia in the East, is

infested with the 31 identified species/subspecies of tsetse. The distribution of different tsetse

groups in the tsetse-infested belt is related to their habitat preferences. The fusca group

species typically occur in the dense, lowland rain forests of West and West-central Africa.

The species belonging to the palpalis group are also basically forest-dwellers, mostly found in

the riverine vegetation of West Africa (Leak, 1999). The species of the morsitans group

typically occur in the savannahs ranging from moist Savannahs or the margins of the forest to

dry savannahs near the margins of the African desert. Figure 2 shows the tsetse belt as well as

the distribution of each of those groups.

Only tsetse flies are biological vector of trypanosome i.e. capable of transmitting the

parasites cyclically. However, other biting insects may mechanically transmit trypanosomes

(Desquesnes and Dia, 2003).


5

Tsetse distribution (All species)

Figure 2: Distribution and number of tsetse fly species belonging to the fusca,

morsitans and palpalis groups in Africa (http://ergodd.zoo.ox.ac.uk/livatl2/tsetse.htm;

Leak, 1999)


6

4. Life cycle of trypanosomes

In sub-Saharan Africa, trypanosomes of mammals are cyclically transmitted by tsetse flies

during their feeding on an infected host. However, some trypanosome species such as T.

evansi or T. vivax can be transmitted mechanically by Tabanids, Stomoxys spp. or other biting

flies (Hoare, 1972). So far known, only T. equiperdum is transmitted sexually. In the case of

cyclical transmission, the parasite has to undergo undergo a developmental cycle within the

insect vector undergoing substantial morphological, biochemical and physiological

transformations to the final metacylic stage that is infective for a new mammalian host. Once

inoculated by tsetse fly bite, these metacyclic trypanosomes undergo development and

multiplication at the site of infection causing a swelling (chancre) and, eventually,

trypomastigotes are released into the blood circulation via the lymphatic system. The parasites

constantly change their surface antigenic coat, the variable surface glycoprotein (VSG),

during their development in the mammal host to escape its immune system.

In the insect vector, after ingesting trypanosomes-infected blood meal, the parasites

undergo a simple or complex cycle of development that takes from a few days to a few weeks

depending on the trypanosome species. During this development, trypanosomes of the

subgenera Trypanozoon and Nannomonas change their morphology and metabolism to

survive in the tsetse’s midgut. This transformation is necessitated by drastic changes in the

parasite’s living conditions i.e. from a stable temperature and oxygen-rich environment in the

vertebrate bloodstream where glucose is used for energy to a fluctuating temperature and

oxygen deficient environment in the tsetse where proline becomes their source of energy

(Vickerman, 1985; Vickerman et al., 1988; Leak, 1999).

Trypanosoma vivax has the simplest life cycle in the tsetse fly, normally developing in

the proboscis (Gardner, 1989). Its development is complete within a period of 5-13 days

(Leak, 1999). Trypanosoma congolense develops first in the midgut and then in the proboscis

of the tsetse fly. Its developmental cycle is longer than that of T. vivax (about 14 days, but

variable ranging from 7- 40 days) (Dale et al., 1995). Trypanosoma brucei spp. establish as

the procyclic form in the fly’s midgut before maturing into the metacyclic trypomastigote in

the vector’s salivary glands. Full development of these species takes about 30 days but may

vary between 17-45 days (Hoare, 1970).


7

5. Impact of trypanosomiasis on animal and human health

The impact of trypanosome infection on animal and human health depends on various factors

among which the susceptibility of the host and the pathogenicity of the trypanosome species

involved in the infection. Wild animals usually do not express severe clinical signs but can

carry trypanosome species including T. b. rhodesiense and constitute an important reservoir of

trypanosomes. In susceptible domestic animals, the disease may be acute and highly

devastating but chronic infections are more common. The host-parasite interaction produces

extensive pathology and severe anaemia. Clinically affected animals lose condition and

become weak and unproductive (FAO, 1998). Trypanosomiasis is often fatal and, at the herd

level, its impact is ranging widely. All aspects of production are deprived: fertility is

impaired; milk yields, growth and work output are reduced; and the mortality rate may reduce

herd size (Connor, 1994). Mortelmans (1984) estimated that if trypanosomosis did not exist in

some Africa regions, these areas could carry three to five times more livestock.

In humans, trypanosomiasis occurs in two stages (Priotto and Sevcsik, 2008). The

stage 1 (the haemolymphatic phase) includes non-specific symptoms like headaches and bouts

of fever. This phase generally goes undiagnosed without active sleeping sickness surveillance.

Figure 3: Life cycle of Trypanosoma brucei (source: http://www.who.int/tdr/diseases/

tryp/lifecycle/htm).


8

The stage 2, the later, neurologic phase occurs when the parasite crosses the blood-brain

barrier and can leads to serious sleep cycle disruptions, paralysis, and progressive mental

deterioration, with behaviourial changes, and, ultimately results in death without effective

treatment.

6. Trypanosomiasis control

In tsetse-infested areas, several strategies can be used to control animal trypanosomiasis

(Cuisance et al., 1994). These are mainly orientated towards the elimination of the parasites

from the blood and the prevention of tsetse bite exposure through vector control.

6.1. Control of the parasite in the host

This control is based on the use of trypanocidal drugs. Depending on the control strategy,

drugs are used for curative or preventive purposes. In animal trypanosomiasis, several

compounds among which diminazene aceturate, isometamidium chloride have been used to

combat the parasite. Isometamidium chloride has a prophylactic activity whilst diminazene

aceturate has a short-term therapeutic activity. However, there has been widespread incorrect

use of these drugs which has lead to the development of drug resistance by the parasite

(Geerts et al., 2001). In the case of sleeping sickness, the most widely used drugs to treat the

patients are Suramin, Melarsoprol and Eflornithine (α difluoromethylornithine) depending on

the trypanosome species and the clinical stage of disease. A new treatment using a

combination of Nifurtimox-Eflornithine is available for sleeping sickness (Priotto and

Sevcsik, 2008; WHO and DNDi, 2009). However, up to now, there is no vaccine available to

control human or animal trypanosomiasis.

6.2. Control of the vector

Initially, attempts to control tsetse flies were based on strategies such as game elimination, the

creation of fly barriers to prevent the advance of the vector and widespread bush clearing to

destroy the habitats for breeding and survival. Current vector control interventions involved

the use of insecticides through the sequential aerosol spraying technique, insecticide-treated

targets or insecticide-treated animals (Vale et al., 1988; Bauer et al., 1995; Hargrove et al.,

2000). Another method currently employed to control tsetse flies is the use of highly efficient

fly traps and screens impregnated with synthetic pyrethroid insecticides (Hargrove et al,.

1995). Moreover, the Sterile Insect Technique (SIT) has successfully eradicated tsetse flies

from the island of Zanzibar (Vreysen et al., 2000), but the effectiveness of this technique


9

largely depends on the isolation of the area, the level of reinvasion from neighbouring

populations. Recently, Aksoy et al. (2003) has shown a potential for the development of novel

control strategies aiming to inhibit the tsetse’s capacity to transmit the parasite. One of these

approaches, transgenesis, would involve blocking the vectorial capacity of the fly by

synthesizing parasite inhibitory molecules by symbiotic bacteria the flies harbour in their

midgut.

Apart from the above methods, the use of trypanotolerant animal (naturally tolerant to

the disease) is also an alternative of trypanosomiasis control.

Despite the arsenal of control tools used since many years, the disease persists and

continues to cause dramatic losses in livestock and burden in public health. The persistence of

the disease could be related to several factors among which the complexity of the interaction

between the parasite and its insect vector tsetse fly. Parameters affecting this tsetse-

trypanosome interaction will be detailed in the following chapter.

Chapter 1

Factors affecting the tsetse’s susceptibility to

trypanosome infection: A literature review

Chapter 1: Factors affecting tsetse-trypanosome interaction: Literature review

11

1.1. Establishment and migration/maturation of trypanosome infection

Trypanosome development involves two key stages in tsetse fly: the initial establishment of

the infection in the midgut followed by a maturation phase in which the procyclic parasites

migrate to the mouthparts (for T. congolense) or salivary glands (for T. brucei spp.) and

finally give rise to metacyclic forms that can be transmitted to a new mammalian host.

In the case of T. brucei species, an essential intermediate developmental stage between

midgut and salivary glands takes place in the proventriculus/foregut and proboscis of the fly

(Van Den Abbeele et al. 1999). The complex development of T. brucei sp. is illustrated in

Figure 1.1. To survive in the tsetse midgut, mammalian trypanosomes must radically

transform their metabolism and begin to multiply. Within two to four hours, viable

trypanosomes that express a new surface coat (procyclin) become visible in the midgut as

they start to transform and divide exponentially (Van Den Abbeele et al., 1999; Gibson and

Bailey, 2003). At around three days, a process of attrition is evident leading to the elimination

of the infection in a large proportion of the flies (Gibson and Bailey, 2003). In successful

infections, trypanosomes invade the ectoperitrophic space by contouring or directly

penetrating the peritrophic membrane where they actively divide. From here they move to the

proventriculus where they cease to divide (Van Den Abbeele et al., 1999). After passing

through the proventriculus, mesocyclic trypanosomes reinvade the endoperitrophic space

(Vickerman, 1985). The migrating parasites undergo maturation into metacyclic forms in

salivary glands where they re-acquire the variable antigen coat to become mature infectious

metacyclics (Vickerman, 1985; Van Den Abbeele et al., 1999).

The ability of a tsetse fly to establish a trypanosome infection in the midgut to develop

an infectious metacyclic infection and to transmit it to a mammal host is known as its

vectorial capacity. Le Ray (1989) referred to the tsetse fly Intrinsic Vectorial Capacity (IVC)

as the intrinsic capability of the fly to produce a metacyclic infection. He proposed the IVC of

a given fly population to be the product of the midgut colonisation and migration +

maturation in proboscis (T. congolense) or salivary glands (T. brucei spp.):

IVC = p × m,

where p is the proportion of tsetse allowing bloodstream trypanosomes to establish as

procyclic forms in the midgut (p = n procyclic flies/n fed flies), and m is the proportion of

tsetse flies infected with procyclic trypanosomes that allowed the trypanosomes to migrate


12

and to mature to the metacyclic stage in the proboscis or salivary glands. (m = n metacyclic

infected flies/n procyclic infected flies).

Both establishment (p) and migration/maturation (m) of trypanosomes in tsetse flies

are developmental barriers, reflected by, in most cases, a low frequency of trypanosome

midgut infection following the ingestion of an infected blood meal, and the fact that only a

variable proportion of these midgut infections continue to mature as an infection in the

mouthparts or salivary glands. Moreover, the prevalence of mature trypanosome infections in

tsetse field populations is surprisingly low. For example, the prevalence of different strains of

T. congolense varied between 1.6 - 4.7% in field tsetse G. pallidipes caught in the

southeastern Zambia (Mekata et al., 2008). Even in highly endemic or epidemic areas of T.

brucei infections, the salivary glands infection rates generally fall within the 0.1 to 0.6%

range (Onyango et al., 1971).

Mammal

Salivary glands

Midgut

Proventriculus & foregut

Tsetse fly

Figure 1.1: Diagram illustrating different key stages of Trypanosoma brucei development in

the tsetse fly and the mammal host (Adapted from Peacock et al., 2007)


13

The developmental barriers for trypanosome development within the tsetse fly have been

attributed to a variety of tsetse fly-related factors (endogenous) that interact with the parasite

(Figure 1.2). Moreover, exogenous factors have been shown to affect trypanosome

development in the tsetse fly.

Figure 1.2: Tsetse fly –related and exogenous factors interfering with the tsetse-trypanosome interaction

Trypanosome development in tsetse fly

Midgut establishment (p) Migration / maturation (m)

SYMBIONTS

MOLECULAR INTERACTION

Lectins

Midgut Proteases

Immune system

• Anti-microbial peptides

• Reactive Oxygen species

• EP- midgut protein

• Temperature

• Host blood

• Starvation

• Trypanosome

genotype

Endogenous factors Exogenous factors

PHYSICAL BARRIER


14

1.2. Endogenous factors affecting tsetse-trypanosome interaction

1.2.1. Molecular base interfering with trypanosome infection in tsetse

1.2.1.1. Role of lectins

Lectins constitute a group of proteins or glycoproteins with a range of properties among

which that of binding specific carbohydrates causing agglutination. Midgut lectins with

molecular mass of 26 and 29 kDa have been identified in different tsetse species (Abubakar et

al., 1995; Osir et al., 1995). The role of lectins in tsetse-trypanosome interaction was first

demonstrated by Maudlin and Welburn (1987) and Welburn et al. (1989) who showed that

feeding tsetse flies with specific sugars (D-glucosamine) enhanced midgut infection rate.

Based on this experiment they suggested that lectins in the tsetse midgut are secreted and

interact with trypanosome development. Moreover, Welburn et al. (1994) suggested that the

midgut lectins were responsible for the agglutination of trypanosomes in the fly midgut by

binding to the procyclic surface coat, prior to the trypanosome’s establishment in the tsetse’s

midgut. More recently, a gene that encodes a proteolytic lectin has been isolated from G. f.

fuscipes (Abubakar et al., 2006) and it was suggested to interfere with trypanosome

establishment in tsetse. Despite all these evidences, supporting data for the direct role of

lectins in the tsetse-trypanosome interaction are still limited since a trypanocidal lectin has not

yet been purified from tsetse.

In addition to the interference with the trypanosomes’ establishment, lectins may also

be involved in initiating trypanosome maturation. This was suggested by laboratory studies

which demonstrated that: i) longer exposure to midgut lectins increased the frequency of

maturation and ii) when lectin activity in the midgut was inhibited, the trypanosomes

remained as procyclic forms but when this inhibition was removed maturation was able to

proceed (Welburn and Maudlin, 1989). However, the mechanism by which lectins promote

the maturation process is not well understood.

It has been shown that addition of antioxidants such as ascorbic acid, uric acid,

glutathione and N-acetyl-cysteine to the infective bloodmeal of tsetse flies significantly

increased trypanosome midgut infection rates suggesting that reactive oxygen species (ROS)

promote trypanosome death in the fly midgut (MacLeod et al., 2007a). Moreover, Xing et al.

(2005) demonstrated that glucosamine can scavenge ROS and this can offer an alternative

explanation for the observed increases in midgut infection rates previousely thought to be

linked to the inhibition of trypanocidal lectins by glucosamine (Maudlin and Welburn, 1987).


15

1.2.1.2 Role of tsetse EP- proteins

The midgut of tsetse flies produces a protein called tsetseEP that shows remarkable sequence

identity to the EP form of the procyclin surface coat molecules of midgut established T .b.

brucei trypanosomes. This tsetseEP protein is strongly upregulated after bacterial challenge

(Haines et al., 2005), suggesting that it plays an important role as a component of the immune

system in tsetse. However, the role of this protein in trypanosome development in tsetse flies

remains unknown and is still under investigation.

1.2.1.3. Role of tsetse flies midgut proteases

When blood stream trypanosomes are taken up by the tsetse fly through a blood meal, they are

exposed to a physiologically hostile environment. Induced by the blood meal, various

proteolytic digestive enzymes are actively secreted in the posterior midgut (Cheeseman and

Gooding, 1985). The role of these proteases in tsetse-trypanosome interactions at this

developmental stage remains poorly understood. However, it is assumed that their activity

could be one of the factors interfering with the killing of the slender bloodstream

trypanosomes. Moreover, they apparently promote the transformation of stumpy forms into

procyclic forms (Imbuga et al., 1992; Sbicego et al., 1999) by a mechanism that remains

unknown. Similar observations have been made in other blood-feeding insect vectors. For

example, in Aedes aegypti, trypsin-like proteases are responsible for the destruction of

ingested ookinetes of Plasmodium (Gass and Yeates, 1979). In contrast, once trypanosomes

have established in the tsetse midgut, the procyclic trypomastigotes with EP-procyclin coat on

their surface, are protected against the detrimental action of the tsetse midgut proteases.

1.2.1.4. Role of tsetse immune system

Most of the knowledge on the innate immune system of insects and its interaction with

parasites has been derived from studies with Drosophila melanogaster and Anopheles

gambiae. Insects do not have the antigen-antibody complexes characteristic for the adaptive

immunity of vertebrates but their immune response involves both cellular and humoral

defense mechanisms that are triggered by pattern recognition receptors (PPRs) capable of

specific binding to pathogen associated molecular patterns (PAMPs) (Medzhitov and

Janeway, 2002). These PRRs can mediate microbial killing directly, through phagocytosis, or

indirectly by the triggering of protease cascades leading to defence reactions such as

encapsulation and melanisation (Lehane et al., 2004) or initiate intracellular immune

signalling pathways which regulate the transcription of antimicrobial peptides (AMPs)

(Reviewed in Dimopoulos, 2003).


16

Two main signalling pathways control the insect immune system: the Toll and the

Immunodeficiency (IMD) pathways (Figure 1.3). The molecular components of each pathway

and mechanisms leading to AMP transcription are reviewed by Lehane et al. (2004) and

Bangham et al. (2006) and are shown in Figure 1.3. Gram-positive bacteria and fungi trigger

the activation of the Toll pathway. Peptidoglycan recognition proteins (PGRP) and gram-

negative binding protein (GNBP) recognize the presence of Gram-positive bacteria and fungi

and, through Spaetzle and Toll, activate a proteolytic cascade. This results in the proteolytic

degradation of inhibitor kB (IkB) protein Cactus and activation of the NF-kB proteins Dif and

Dorsal, resulting in the transcription of AMPs (Figure 1.3). The lipopolysaccharides (LPS)

present on Gram-negative bacteria trigger the IMD pathway, which also results in a

proteolytic cascade. This results in the cleavage of Relish (another NF-kB transcription factor)

and activation of AMPs transcription (Figure 1.3) (Reviewed in Lehane et al., 2004;

Boulanger et al., 2006; Bangham et al., 2006). Both the Toll and the IMD pathways regulate

AMPs transcriptions. Some AMPs are specific to one pathway, and others are activated by

both, but little is known about how this specificity is translated into gene-expression profile.

However, Bulet and Stöcklin (2005) suggested that most of the AMPs such as diptericin,

drosocin, cecropins and attacins are controlled by IMD pathway while Hoffmann et al. (1996)

concluded that AMPs cecropins, attacins and defensins appear to be controlled by both

pathways.

Most insect AMPs are composed of 20-40 amino acids and their primary structures

adopt an α-helical conformation. Some AMP sequences contain pairs of cysteine residues

(Dimopoulos, 2003; Bulet and Stöcklin, 2005). In dipterian insects, AMPs are synthesized

principally by the fat body and released into the hemolymph, but they can also be expressed

by the hemocytes and various epithelia, particularly the anterior part of the gut (Lehane et al.,

1997). The activity spectrum of immune peptides is in most cases specific for different classes

of pathogens and the bacteria killing mechanism is believed to rely on the disintegration of the

bacterial membrane or the interference with membrane assembly or bacterial proteins (Otvos,

2000).


17

The tsetse fly immune system has been found to play a role in determining the

efficiency of trypanosome establishment (Hao et al., 2001). Indeed, infection rates in the

tsetse midgut decreased significantly when the teneral tsetse fly immune response was

artificially stimulated by Escherichia coli or LPS prior to the infectious blood meal (Hao et

al., 2001). This indicated that immune events early in the infection process are apparently

affecting the trypanosome developmental outcome in the fly. Moreover, laboratory

investigations with several immune marker genes suggested that the fat body, the midgut and

proventriculus tissue, all have a role in the tsetse immune response. To understand the role of

the fat body response during trypanosome transmission, Hao et al. (2001) studied the

transcriptional regulation of the immune genes attacin, defensin and diptericin in vivo during

the natural process of the parasite infection. These studies showed that upon entering the

Figure 1.3: The molecular components involved in the two major signalling pathways in the

Drosophila immune response (Adapted from Bangham et al., 2006).

Abbreviations: PGRP (peptidoglycan-recognition proteins; GNBP (gram negative binding

proteins)


18

tsetse via the blood meal, trypanosomes failed to elicit a strong immune response in any of the

immune tissues, contrary to what is observed in response to E. coli. Similarly, when

trypanosomes were injected into the hemolymph, no immediate stimulation of the systemic

responses was observed. Similar injection of bacteria (E. coli), on the other hand, resulted in

abundant response. In contrast, when the same immune genes were analysed in older flies

with established trypanosome infection in the midgut, immune genes were observed in the

hemolymph (Boulanger et al., 2002) and in the fat body and proventriculus tissues (Hao et al.,

2003). All these findings suggested that the bloodstream form trypanosomes, present in the

early phase of infection, do not induce immune responses in the tsetse fly. However, after

differentiation into the procyclic forms, trypanosomes are recognized as foreign and elicit

responses in the fat body as well as in the proventriculus. Apparently, the energy invested by

tsetse to respond immunologically to trypanosome infection results in a significant reduced of

their fecundity (Hu et al., 2008) suggesting that the parasite infection has negative effects on

the flies. Because trypanosomes remain in the midgut without crossing into the hemolymph, it

is possible that reactive intermediates such as nitric oxide (NO) or hydrogen peroxide (H2O2)

function as immunological signals mediating molecular communication between the different

tsetse compartments (Hao et al., 2003). Implication of oxidative stress in immune defenses

has been invoked in Drosophila (Ha et al., 2005) and more recently in tsetse flies (MacLeod

et al., 2007b).

As the response elicited late in the infectious process fails to clear the parasite

infections, trypanosomes might either be resistant to actions of the immune peptides, though

trypanolytic activity has been demonstrated for tsetse Attacin peptide (Hu and Aksoy, 2005)

or they fail to reach and harm the parasites in their midgut niche.

In addition to the pathogen-specificity of the tsetse’s immune response, capable of

discriminating not only between bacteria and trypanosome but also between bloodstream form

and procyclic trypanosomes, the immune response has been correlated to tsetse’s

susceptibility to trypanosome infection. In this respect, susceptible flies are expected to have a

higher level of immune gene expression in the proventriculus and midgut than non-

susceptible flies (Nayduch and Aksoy, 2007). Except for attacin (Hu and Aksoy, 2005), the

underlying mechanisms by which the tsetse’s antimicrobial peptides interfere with

trypanosome development remain largely unknown and need further investigation.


19

1.2.2. Role of endosymbionts in the development of a trypanosome infection

Tsetse flies harbor multiple symbiotic microbes among which two members of the

Enterobacteriaceae i.e. the obligate mutualist Wigglesworthia and the commensal mutualist

Sodalis glossinidius (Aksoy, 1995). Wigglesworthia resides within differentiated midgut cells

whereas Sodalis glossinidius is present in different tissues such as midgut, hemolymph and

salivary glands (Cheng and Aksoy, 1999). These symbionts influence several important

physiological events in the flies. Indeed, their elimination through the application of

antibiotics was shown to have a dramatic impact on tsetse fecundity and pupal emergence,

effectively rendering the insect sterile (Dale and Welburn, 2001). Moreover, elimination of

Sodalis glossinidius resulted in increased refractoriness to T. b. rhodesiense infection in

tsetse, suggesting that this symbiont plays a role in potentiating susceptibility to trypanosome

infection in tsetse (Dale and Welburn, 2001). Geiger et al. (2005, 2007) suggested that the

presence of specific genotypes of Sodalis glossinidius may be related to the different vector

competences of Glossina spp. The mechanism by which symbionts potentiate midgut

infection remains largely unknown. Wang et al. (2009) recently showed that a tsetse pathogen

recognition protein is synthesized a in the bacteriome organ in response to the presence of

Wigglesworthia and prevents the activation of tsetse’s immune responses, which may

influence the fly’s ability to trypanosome transmission.

1.3. Exogenous factors affecting tsetse-trypanosome interaction

1.3.1. Temperature

Several laboratory studies positively correlated increased susceptibility with the maintenance

temperature of puparia or adult flies. Indeed, Kinghorn and Yorke (reviewed in Leak, 1999)

found that T. b. rhodesiense developed more readily in G. morsitans kept at higher

temperature. Similarly, G. morsitans flies emerging from pupae incubated at 30°C were

shown to develop high infection rates with T. b. brucei (Burtt, 1946) or T. congolense

(Ndegwa et al., 1992). Under field conditions, Ford and Leggate (1961) have found a

correlation between high infection rates and increased mean annual temperature, although

other factors may play a role in an uncontrolled natural environment.

The effect of tsetse cooling on trypanosome development is controversial Chilling of

tsetse flies to 0-5°C for 30 min post-infection was shown to increase midgut infections but did

not affect significantly the proportion of trypanosome infections maturing in male tsetse flies

(Otieno et al., 1983). In contrast, when Macleod et al. (2007b) chilled tsetse flies for a short


20

period, midgut infection rates were not affected but this chilling boosted maturation of

trypanosome infection in female flies but not in males.

The underlying mechanism by which temperature interferes with tsetse-trypanosome

interactions is not clear. It has been shown that chilling induced the synthesis of heat shock

proteins in Drosophila (Burton et al., 1988) and the effect of chilling in tsetse may probably

follow similar patterns.

1.3.2. Role of host blood

Differences in midgut infection and its subsequent maturation to metacyclic trypanosome

have been found when various mammalian hosts’ blood has been used for the infective meal.

For example, buffalo and eland blood supported low midgut infection rates in the tsetse fly

compared to goat blood which enhances infections (Mihok et al., 1993; 1995). Similarly,

maintaining tsetse flies on defibrinated bovine blood resulted in a significant reduced

proportion of flies that developed a trypanosome infection (Zongo et al., 2004). The ability of

blood to facilitate the transformation process from bloodstream form to procyclic

trypanosomes appears crucial for successful establishment of infections in tsetse (Nguu et al.,

1996). The mechanisms by which host blood modulates midgut infection in tsetse are still not

clearly understood. It may be due to the composition of the blood. Serum, for example, is

thought to stimulate the secretion of digestive enzymes but also to trigger the secretion of

trypanocidal lectins in the midgut. Since removal of serum from tsetse diet was shown to

inhibit maturation (Maudlin et al., 1984), the serum components may probably play a primary

role enabling the procyclic midgut trypanosome to mature. More studies are required to

further elucidate the phenomenon.

1.3.3. Role of tsetse fly starvation

Evidence on the effect of starvation on the fly’s subsequent vector competence is sparse and

sometime contradicting. While Gooding (1988) did not find significant differences in the

prevalence of T. b. brucei infections in G. m. centralis starved for five days after the infective

feed, Gingrich et al. (1982) observed a significant increased proportion of four-day starved

older G. m. morsitans that developed a metacyclic infection with T. b. rhodesiense. More

recent experiments showed unequivocally that starvation enhanced the tsetse’s susceptibility

to trypanosome infections. Indeed, starving teneral and 20-days-old tsetse flies for several

consecutive days resulted in significant increases in T. congolense as well as in T. b. brucei

infections (Kubi et al., 2006).


21

The underlying mechanisms that cause this important change in susceptibility of the

vector after starvation still need to be clarified. However, it has been hypothesized that the

exposure of the flies to a nutritional stress up to a level that causes a depletion of its energy

reserve may result in down-regulation of immune system and subsequently leads to a less

hostile midgut environment for ingested trypanosomes. Indeed, the fat body constitutes the

tsetse’ energy reserve and functions as a key centre of metabolism and biochemistry control

for the fly’s innate immune response. This association between starvation, fat depletion and

immunological responses still requires further investigations.

1.3.4. Role of trypanosome genotype

Apart from these tsetse-related factors described above, susceptibility of tsetse flies to a

trypanosome infection can also be influenced by the parasite itself. Results from various field

and experimental studies revealed that, in most cases, high infection rates are generally

observed with T. congolense rather than T. brucei (Woolhouse et al., 1994; Mohamed-

Ahmeed et al., 1989). Moreover, significant variations have also been reported within a

trypanosome species. For example, T. b. brucei produces higher midgut and salivary glands

infections in tsetse flies compared to T. b. rhodesiense (Welburn et al., 1995). In addition,

variations in procyclic and metacyclic infection rates have been observed as a result of

changes in some biological properties of the parasite such as the increased resistance to

trypanocidal drugs that is associated with an increased transmissibility (Van den Bossche et

al., 2006). In the same context, T. congolense strains of high virulence profile were seemed to

exhibit high potential of transmissibility by tsetse flies compared to trypanosome strains of

moderate or low virulence profile (Masumu et al., 2006).

1.4. Conclusion

The developmental cycle of trypanosome within tsetse fly is complex and is mediated by a

variety of factors. With regard to the tsetse-related factors, the nutritional status seems to play

an important role. Through starvation adult flies regain a higher susceptibility to infection

which contradicts with the generally accepted trypanosome transmission paradigm. It thus

seems that nutritional stress and associated starvation changes the tsetse’s susceptibility to

infection with pathogenic trypanosomes species and puts the epidemiology of trypanosome in

a new perspective. Nevertheless, some questions such as the transmission rate of human

infective trypanosomes by starved flies, impact of starvation of reproducing female tsetse flies

on their offspring’s susceptibility to trypanosomal infections and the progress of established

midgut infection in nutritionally stressed flies, need to be clarified. Moreover, the underlying


22

physiological mechanism(s) that induce(s) the change in this susceptibility remains largely

unknown and need further investigations. It is difficult to study the impact of starvation on

each tsetse’s intrinsic component interfering with trypanosome infection. Therefore, we

focused our studies on the immune system with the hypothesis that starvation affects the

tsetse immune system and subsequently reduces its immune response to pathogen challenge

including trypanosomes. It is expected that elucidating all these aspects may contribute to a

better understanding of the epidemiology of tsetse-transmitted trypanosomiasis and to the

development of appropriate control strategies.


23

1.5. References

Abubakar, L.U., Bulimo, W.D., Mulaa, F.J., Osir, E.O., 2006. Molecular characterization of a tsetse fly midgut proteolytic lectin that mediates differentiation of African trypanosomes. Insect Biochemistry and Molecular Biology 36: 344-352

Abubakar, L., Osir, E.O., Imbuga, M.O., 1995. Properties of a bloodmeal-induced midgut lectin from the tsetse fly Glossina morsitans. Parasitology Research 81: 271-275

Aksoy, S., Gibson, W.C., Lehane, M.J., 2003. Interactions between tsetse trypanosomes with implications for the control of trypanosomiasis. Advances in Parasitology 53:1-83

Aksoy, S., 1995. Wigglesworthia gen. nov. and Wigglesworthia glossinidia sp. Nov., Taxa Consisting of the Mycetocyte-Associated, Primary Endosymbionts of Tsetse Flies. International Journal of Systematic Bacteriology 45: 848-851

Bangham, J., Jiggins, F., Lemaitre, B., 2006. Insect Immunity : The Post-Genomic Era. Immunity 25: 1-5

Barrett, M.P., 1999. The fall and rise of sleeping sickness. Lancet 353: 1113-1114

Bauer, B., Amsler-Delafosse, S., Clausen, P.H., Kabore, I., Petrich-Bauer, J., 1995. Successful application of deltamethrin pour-on to cattle in a campaign against tsetse flies (Glossina spp) in the pastoral zone of Samorogouan, Burkina Faso. Tropical

Medicine and Parasitology 46: 183-189

Boulanger, N., Bulet, P., Lowenberger, C., 2006. Antimicrobial peptides in the interactions between insects and flagellate parasites. Trends in Parasitology 22 : 262-268

Boulanger, N., R. Brun, L. Ehret-Sabatier, C. Kunz, P., Bulet, P., 2002. Immunopeptides in the defense reactions of Glossina morsitans to bacterial and Trypanosoma brucei brucei

infections. Insect Biochemistry and Molecular Biology 32: 369-375.

Bulet, P., Stöcklin, R., 2005. Insect Antimicrobial Peptides: Structures, Properties and Gene Regulation. Protein and Peptides Letters 12: 3-11

Burton, V., Mitchell, H.K., Young, P., Petersen N.S., 1988. Heat shock protection against cold stress of Drosophila melanogaster. Molecular and Cell Biology 8: 3550-3552.

Burtt, E.D., 1946. Incubation of tsetse pupae: increased transmission rate of Trypanosoma

rhodesiense in Glossina morsitans. Annale of Tropical Medicine and Parasitology 40:18-28

Buxton, P.A., 1955. The natural history of tsetse flies. An account of the biology of the genus Glossina (Diptera). Lewis H.K. & Co Ltd, London, 816 pp.

Cheeseman, M.T., Gooding, R.H., 1985. Proteolytic enzymes from the tsetse flies Glossina

morsitans and Glossina palpalis (Diptera : Glossinidae). Insect Biochemistry 15: 677-680


24

Cheng, Q., Aksoy, S., 1999. Tissue tropism, transmission and expression of foreign genes in vivo in midgut symbionts of tsetse flies. Insect Molecular Biology 8:125-132

Connor, R.J., 1994. The impact of nagana. Onderstepoort Journal of Veterinary Research 61: 379-383.

Cuisance, D., Barre, N., De Deken, R., 1994. Ectoparasites des animaux : méthodes de lutte écologique, biologique, génétique et mécanique. Revue Scientifique et Technique de

l’Office International des Epizooties 13 : 1305-1356

Dale, C., Welburn, S. C., Maudlin, I. and Milligan, P. J., 1995. The kinetics of maturation of trypanosome infections in tsetse. Parasitology 111: 187-191

Dale, C., Welburn, S.C., 2001. The endosymbionts of tsetse flies: manipulating host-parasite interactions. International Journal for Parasitology 31: 628-631

Desquesnes, M., Dia, M.L., 2003. Mechanical transmission of Trypanosoma congolense in cattle by the African tabanid Atylotus agrestis. Experimental Parasitology 105: 226-231.

DFID, 2001. Trypanosomosis, Tsetse and Africa. The year 2001 report, 15 pp.

Dimopoulos, G., 2003. Insect immunity and its implication in mosquito-malaria interactions. Cellular Microbiology 5: 3-14

Food and Agriculture Organization (FAO), 1998. A field guide for the diagnosis, treatment and prevention of African animal trypanosomosis by G. Uilenberg. FAO, Rome: 158pp

Ford, J., Leggate, B.M., 1961. The geographical and climatic distribution of trypanosome infection rates in Glossina morsitans group of tsetse-flies. Transactions of the Royal

Society for Tropical Medicine and Hygiene 55: 383-397

Gardner, P.R., 1989. Recent studies of the biology of Trypanosoma vivax. Advances in

Parasitology 28: 229-317

Gass, R.F., Yeates, R.A., 1979. In vitro damage of cultured ookinetes of Plasmodium gallinaceum by digestive proteinases from susceptible Aedes aegypti. Acta Tropica 36: 243-252

Geiger, A., Ravel, S., Mateille, T., Janelle, J., Patrel, D., Cuny, G., Frutos, R., 2007. Vector Competence of Glossina palpalis gambiensis for Trypanosoma brucei s.l. and Genetic Diversity of the Symbiont Sodalis glossinidius. Molecular Biology Evolution 24: 102-109

Geiger, A., Ravel, S., Frutos, R., Cuny, G., 2005. Sodalis glossinidius (Enterobacteriaceae) and Vectorial Competence of Glossina palpalis gambiensis and Glossina morsitans morsiatns for Trypanosoma congolense Savannah Type. Current Microbiology 51: 35-40

Geerts, S., Holmes, P.H., Diall, O., Eisler, M.C., 2001. African bovine trypanosomiasis: the problem of drug resistance. Trends in Parasitology 17: 25-28


25

Gibson, W., Bailey, M., 2003. The development of Trypanosoma brucei within the tsetse fly midgut observed using green fluorescent trypanosomes. Kinetoplastid Biology and

Disease, www.kinetoplastids.com/content/2/1/1

Gingrich, J.B., Ward, R.A., Macken, L.M., Esser, K.M., 1982. African sleeping sickness: new evidence that mature tsetse flies (Glossina morsitans) can become potent vector. Transactions of the Royal Society of Tropical Medicine and Hygiene 76: 479-481

Gooding, R., 1988. Infection of post teneral tsetse flies (Glossina morsitans morsitans and Glossina morsitans centralis) with Trypanosoma brucei brucei. Canadian Journal of

Zoology 66: 1289-1292

Ha, E.M., Oh, C.T., Bae, Y.S., Lee, W.J., 2005. A direct role for dual oxidase in Drosophila gut immunity. Science 310: 847-850

Haines, L.R., Jackson, A.M., Lehane, M.J., Thomas, J.M., Yamaguchi, A.Y., Haddow, J.D., Pearson, T.W., 2005. Increased expression of unusual EP repeat-containing proteins in the midgut of the tsetse fly (Glossina) after bacterial challenge. Insect Bioschemistry

and Molecular Biology 35: 413-423

Hao, Z., Kasumba, I., Aksoy, S., 2003 Proventriculus (cardia) plays a crucial role in immunity in tsetse fly (Diptera: Glossinidiae). Insect Biochemistry and Molecular Biology 33: 1155-1164

Hao, Z., Kasumba, I., Lehane, M.J., Gibson, W.C., Kwon, J., Aksoy, S., 2001. Tsetse immune responses and trypanosome transmission: implications for the development of tsetse-based strategies to reduce trypanosomiasis. Proceedings of the National Academy of

Sciences of the USA 98:12648-12653

Hargrove, J.W., Silas, O., Msalilwa,J.S.I., Fox, B. 2000. Insecticide-treated cattle for tsetse control: The power and the problems. Medical and Veterinary Entomology 14: 123-130

Hargrove, J.W., Holloway, M.T.P., Vale, G.A., Gough, A.J.E., Hall, D.R., 1995. Catches of tsetse (Glossina spp.) (Diptera Glossinidae) from traps and targets baited with large doses of natural and synthetic host odour. Bulletin of Entomological Research 85: 215-227

Hoare, C. A., 1972. The trypanosomes of Mammals. A Zoological Monograph. Blackwell, Oxford, 749 pp.

Hoare, C.A. , 1970. The mammalian trypanosomes of Africa. In: Mulligan, H.W. (ed.) The African trypanosiamiases. George Allen and Unwin, London, pp.3-23.

Hoffmann, J.A., Jean-Marc Reichhart, J.M., Hetru, C., 1996. Innate immunity in higher insects. Current Opinion in Immunology 8: 8-13

Hu, J.W.C., Wu, Y., Stuart, A., Amemiya, C., Berriman, M., Toyoda, A., Hattori, M., Aksoy, S., 2008. Characterization of the antimicrobial peptide attacin loci from Glossina

morsitans. Insect Molecular Biology 17: 293-302


26

Hu, C., Aksoy, S., 2005. An Antimicrobial peptide with trypanocidal activity characterised from characterised from Glossina morsitans morsitans. Insect Biochemistry and

Molecular Biology 35:105-115

Imbuga, M.O., Osir, E.O., Labongo, V.L., Darji, N., Otieno, L.H., 1992. Studies on tsetse midgut factors that induce differentiation of bloodstream Trypanosoma brucei brucei in vitro. Parasitology Resaerch 78: 10-15

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Van den Bossche, P. 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392

Leak, S. A.G., 1999. Tsetse biology and ecology. Their role in the epidemiology and control of trypanosomosis. CABI publishing, UK, 529 pp

Lehane, M.J., Aksoy, S., Levashina, E., 2004. Immune responses and parasite transmission in blood-feeding insects. Trends in Parasitology 20: 433-439

Lehane, M.J, Wu, D., Lehane, S.M., 1997. Midgut-specific immune molecules are produced by the blood-sucking insect Stomoxys calcitrans. Proceedings of the National Academy

of Sciences of the USA 94: 11502–11507

Le Ray, D., 1989. Vector susecptibility to African trypanosomes. Annales de la Société Belge

de Médecine Tropicale 69: 165-171

Macleod, E.T., Maudlin, I., Darby, A.C., Welburn, S.C., 2007a. Antioxidants promote establishment of trypanosome infections in tsetse. Parasitology 134: 827-831

Macleod, E.T., Darby, A.C., Welburn, S.C., 2007b. Factors affecting trypanosome maturation in tsetse flies. PLOS One, 2, e239. doi:10.137/journal.pone.0000239.

Masumu Mulumbu, J., 2006. The importance of Trypanosoma congolense strain diversity in the epidemiology of bovine. PhD Thesis. Universiteit Gent, Gent, 115pp.

Masumu, J., Marcotty, T., Ndeledje, N., Kubi, C., Geerts, S., Vercruysse, J., Dorny, P., Van den Bossche, P., 2006. Comparison of the transmissibility of Trypanosoma congolense strains, isolated in a trypanosomiasis endemic area of eastern Zambia, by Glossina morsitans morsitans. Parasitology 133: 331-334

Meddzhitov, R., Janeway, C.A.J., 2002. Decoding the patterns of self and nonself by innate immune system. Sciences 296: 298-300

Mekata, H., Konnai, S., Simuunza, M., Chembensofu, M., Kano, R., Witola, W.H., Tembo, M.E., Chitambo, H., Inoue, N., Onuma, M., Ohashi, K., 2008. Prevalence and source of trypanosome infections in field-captured vector flies (Glossina pallidipes) in southeastern Zambia. Journal of Veterinary Medical Science 70:923-928

Maudlin, I., Welburn, S.C., 1987. Lectin mediated establishment of midgut infections of Trypanosoma congolense and Trypanosoma brucei in Glossina morsitans. Tropical



27

Maudlin, I., Kabayo, J.P., Flood, M.E.T., Evans, D.A., 1984. Serum factors and the maturation of Trypanosoma congolense infections in Glossina morsiatns. Zeitschrift für

Parasitenkunde 70: 11-19

Mihok, S., Machika, C., Darji, N., Kang'ethe, E.K., Otieno, L.H., 1995. Relationships between host blood factors and proteases in Glossina morsitans subspecies infected with Trypanosoma congolense. Medical and Veterinary Entomology 9: 155-160

Mihok, S., Olubayo, R.O., Darji, N., Zweygarth, E., 1993. The influence of host blood on infection rates in Glossina morsitans spp. infected with Trypanosoma congolense, T. brucei and T. simiae. Parasitology 107: 41-48

Mohamed-Ahmed, M.M., Ahmed, A.I., Ishag, A., 1989. Trypanosome infection rate of Glossina morsitans submorsitans in Bahr El Arab, South Darfur Province, Sudan. Tropical Animal Health and Production 21: 239-244

Mortelmans J. 1984. Socio-Economic problems related to animal trypanosomiasis in Africa. Social Sciences and Medicine 19: 1105-1107

Nayduch, D., Aksoy, S., 2007. Refractoriness in Tsetse Flies (Diptera: Glossinidae) May be a Matter of Timing. Journal of Medical Entomology 44: 660-665

Ndegwa, P.N., Irungu, L.W., Moloo, S.K., 1992. Effect of puparia incubation temperature: increased infection rates of Trypanosoma congolense in Glossina morsitans centralis,

G. fuscipes fuscipes and G. brevipalpis. Medical and Veterinary Entomology 6:127-130

Nguu, E.K., Osir, E.O., Imbuga, M.O., Olembo, N.K., 1996. The effect of host blood in the in

vitro transformation of bloodstream trypanosomes by tsetse midgut. Medical and

Veterinary Entomology 10: 317-322

Onyango, R.J., Geigy, R., Mwanbu, P.M., Moloo, S.K., 1971. The endemicity of Rhodesian sleeping sickness in Ikoma-Serengeti area: final discussion. Acta Tropica 28: 221-225

Otieno, L.H., Darji, N., Onyango, P., Mpanga, E., 1983. Some observations on factors associated with the development of Trypanosoma brucei brucei infection in Glossina morsitans morsitans. Acta Tropica 40: 113-120

Otovos, L. J., 2000. Antibacterial peptides isolated from insects. Journal of Peptide Science 6:497-511

Osir, E.O., Abubakar, L., Imbuga, M.O., 1995. Purification and characterisation of a midgut lectin-trypsin complex from the tsetse fly Glossina longipennis. Parasitology Research 81: 276-281

Peacock, L., Ferris, V., Bailey, M., Gibson, W., 2007. Dynamics of infection and competition between two strains of Trypanosoma brucei brucei in the tsetse fly observed using fluorescent markers. Kinetoplastid Biology and Disease 6: 4 doi:10.1186/1475-9292-6-4

Priotto, G., Sevcsik, A.M., 2008. Nifurtimox-Eflornithine Combination Therapy: An improved treatment for sleeping sickness. The Drugs for Neglected Diseases initiative (DNDi) newsletter 17: 1-8


28

Sbicego, S., Vassella, E., Kurath, U., Blum, B., Roditi, I., 1999. The use of trangenic Trypanosoma brucei to identify compounds inducing the differentiation of bloodstream forms to procyclic forms. Molecular and Biochemical Parasitology 104: 311-322

Soltys, M.A., 1963. Immunity in African trypanosomiasis. Bulletin of the World Health

Organization 28: 753-761

Stephen, L.E., 1986. Trypanosomiasis : A veterinary perspective. Pergamon Press, 551 pp.

Swallow, B.M., 2000. Impacts of trypanosomiasis on African Agriculture. PAAT Technical

and Scientific series (FAO) 2: 52 pp.

Vale, G.A., Lovemor, D.F., Flint, S., Cockbill, G.F., 1988. Odour-baited targets to control tsetse flies, Glossina spp. (Diptera: Glossinidae), in Zimbabwe. Bulletin of

Entomological Research 78: 31-49

Van Den Abbeele, J., Claes, Y., van Bockstaele, D., Le Ray, D., Coosemans, M., 1999. Trypanosoma brucei spp. development in the tsetse fly: characterization of the postmesocyclic stages in the foregut and proboscis. Parasitology 118: 469-478

Van den Bossche, P., Akoda, K., Kubi, C., Marcotty, T., 2006. The transmissibility of Trypanosoma congolense seems to be associated with its level of resistance to isometamidium chloride. Veterinary Parasitology 135: 365-367

Vickerman, K., 1985. Developmental cycles and biology of pathogenic trypanosomes. British

Medical Journal 41:105-114

Vickerman, K., Tetley, L., Hendry, K.A., Turner, C.M., 1988. Biology of African trypanosomes in the tsetse fly. Biology of the Cell 64: 109-119

Vreysen, M.J.B ., Saleh, K.M ., Ali, M.Y ., Abdulla, A.M ., Zhu, Z-R ., Juma, K.G ., Dyck, A.V., Msangi, A.R., Mkonyi, P.A., Feldman, H.U., 2000. Glossina Austeni (Diptera: Glossinidae) eradicated on the island of Unguja, Zanzibar, using the Sterile insect technique. Journal of Economic Entomology 93: 123-135

Wang, J., Wu, Y., Yang, G., Aksoy, S., 2009. Interactions between mutualist Wigglesworthia and tsetse peptidoglycan recognition protein (PGRP-LB) influence trypanosome transmission. Proceedings of the National Academy of Sciences of the USA, 106: 12133-12138

Welburn, S.C., Maudlin, I., 1989. Lectin signalling of maturation of T. congolense infections in tsetse. Medical and Veterinary Entomology 3:141-145

Welburn, S.C., Maudlin, I., Ellis, D.S.,1989. Rate of trypanosome killing by lectins in midguts of different species and strains of Glossina. Medical and Veterinary

Entomology, 3, 77-82

Welburn, S.C., Maudlin, I., Molyneux, D.H., 1994. Midgut lectin activity and sugar specificity in teneral and fed tsetse. Medical and Veterinary Entomology 8: 81-87


29

Welburn, S.C., Maudlin, I., Milligan, P.J.M., 1995. Trypanozoon: infectivity to human is linked to reduced transmissibility in tsetse. I. Comparison of human serum-resistant and human serum-sensitive field isolates. Experimental Parasitology 81: 404-408

Woolhouse, M.E., Bealby, K., McNamara, J.J., Silutongwe, J., 1994. Trypanosome infections of the tsetse fly Glossina pallidipes in the Luangwa Valley, Zambia. International

Journal for Parasitology 24: 987-993

World Health Organization, 2006. Human Africa trypanosomiasis (sleeping sickness). Weekly

Epidemiological Record 81, 71-80

WHO (World Health Organozation) and DNDi (The Drugs for Neglected Diseases initiative), 2009. Progress made and challenges ahead in clinical research and development of new treatment for human African Trypanosomiasis. 30th International Scientific Council for Tsetse Research and Control (ISCTRC) meeting, Kampala, Uganda, 21-25 September

Xing, R., Liu, S., Guo, Z., Yu, H., Li, C., Ji, X., Feng, J., Li, P., 2005. The antioxidant activity of glucosamine hydrochloride in vitro. Bioorganic and Medicinal Chemistry 14: 1706-1709

Zongo, I., Mbahin, N., Van Den Abbeele, J., De Deken, R., Van den Bossche, P., 2004. Comparison of the infection rate of tsetse, Glossina morsitans morsitans, fed in vitro or in

vivo. Medical and Veterinary Entomology 18: 64-66

Objectives of the thesis

Objectives

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The main objective of this thesis was to contribute to our understanding of factors that

influence the ability of the tsetse fly to transmit animal and human pathogenic trypanosomes,

in experimental and in natural conditions.

The specific objectives were:

� To evaluate the susceptibility of nutritionally stressed tsetse flies for infection with the

human pathogenic trypanosomes T. b. rhodesiense and T. b. gambiense;

� To evaluate the effect of nutritional stress on the progress of an established T. brucei

brucei midgut infection to metacyclic trypanosomes in the salivary glands;

� To determine the effect of nutritional stress of reproducing female tsetse flies on the

susceptibility of their offspring to trypanosome infection;

� To determine whether nutritional stress affects the tsetse baseline immune peptides

expression and its immune response to trypanosomal and bacterial challenges;

� To determine the effect of nutritional stress on the tsetse fly immune status and

trypanosome infection in natural, field-caught flies;

� To investigate whether the developmental phase of a monomorphic T. congolense

strain in the mammalian host affects transmissibility with special emphasis on the

nutritional status of the fly.

Chapter 2

Effect of starvation on the tsetse fly’s susceptibility to human

pathogenic trypanosome species (T. b. rhodesiense and T. b. gambiense)

A manuscript based upon the T. b. rhodesiense infection data has been submitted to

‘Emerging Infectious Disease’

Akoda K., Van Den Abbeele J., Van Reet N., De Deken R, Marcotty T., Lejon V.,

Coosemans M., Van den Bossche P.(-) A highly transmissible Trypanosoma brucei

rhodesiense strain recently isolated from a sleeping sickness patient in Malawi.

Chapter 2: Nutritional stress and T.b. rhodesiense, T.b. gambiense transmission

33

2.1. Introduction

Human African trypanosomiasis (HAT) or sleeping sickness is a neglected tropical parasitic

disease, unique to sub-Saharan Africa, which causes a considerable public health burden on

rural populations (Barret, 1999). The disease is caused by two subspecies of Trypanosoma

brucei spp., T. b. gambiense and T. b. rhodesiense, both cyclically transmitted by tsetse flies.

The epidemiology of this important disease is determined largely by the number of infected

flies in a tsetse population. The proportion of infected tsetse is determined by a range of

intrinsic and extrinsic factors. For T. b. rhodesiense, several transmission experiments

showed that the parasite transmissibility is generally low, with a metacyclic infection rate in

laboratory reared or field captured Glossina morsitans morsitans usually below 10% (Burtt,

1946; Harley, 1971; Gingrich et al., 1982a). Welburn et al. (1995) attributed this low

transmissibility of T. b. rhodesiense, compared to that of T. b. brucei, to its infectivity to

humans. Moreover, surprisingly few tsetse infected with T. b. gambiense (0.1-0.5 %) have

been detected in areas where many human cases were nonetheless observed (Jamonneau et al.,

2004). However, environmental factors such as the high ambient temperature have been

shown to increase the tsetse susceptibility to infection with T. b. rhodesiense (Taylor, 1932;

Burtt, 1946; Fairbairn and Watson, 1955; Ndegwa et al., 1992). Recent studies have shown

that the nutritional status of tsetse flies at the time of the infective blood meal also affects the

fly susceptibility to infection with T. congolense or T. b. brucei (Kubi et al., 2006). However,

only few and preliminary experimental data on the effect of starvation on the transmissibility

of human pathogenic trypanosome species in tsetse flies are available (Gingrich et al., 1982b).

To determine the effect of starvation on the susceptibility of tsetse to T. b. gambiense

or T. b. rhodesiense a number of experiments was conducted. To mimic the field situation as

much as possible, use was made of tsetse species belonging to the savannah (G. m. morsitans)

or the riverine subgenera (G. p. gambiensis).

2.2. Material and methods

2.2.1. Trypanosome isolates

The trypanosome strain T. b. rhodesiense RUMPHI was used, a strain isolated in 2007 at the

hospital of Rumphi, Malawi, from a sleeping sickness patient who had relapsed 24 months

after treatment with suramin. After 4 passages in mice, this strain was adapted to HMI-9, with

15% foetal calf serum instead of Serum Plus (Hirumi, 1989) and confirmed to be human

serum-resistant (HSR) when incubated with an extra 5% human serum. Furthermore, this

strain tested PCR positive for the serum resistance associated gene (SRA). In another set of


34

experiments, use was made of the T. b. gambiensis S1/1/6 strain. This trypanosome strain was

isolated in 2002 from a HAT patient in Côte d’Ivoire (Ravel et al., 2006). In addition,

different strains of T. b. gambiense (MBA and NHOM/INRB/2007) available in the cryobank

of the Institute of Tropical Medicine (ITM, Antwerp/Belgium) and T. b. gambiense S12.9.5

obtained from IRD-CIRAD laboratory (Montpellier/France) were used in consecutive

infection experiments.

2.2.2. Tsetse flies and experimental groups

For infections with T. b. rhodesiense, male G. m. morsitans Westwood from the colony

maintained at the ITM were used. The origin of this tsetse colony and the rearing technique

are described by Elsen et al. (1993).

The transmissibility of the T. b. rhodesiense isolate was investigated in six groups of

experimental flies that differed in age and/or in their nutritional status: i) Group 1: non-

starved teneral flies (= newly emerged unfed flies less than 32 hours old, ii) Group 2: teneral

flies that were starved for four days before the infective blood meal (starved tenerals, iii)

Group 4: 20-day-old flies starved for two days after the last blood meal (non-starved old flies

and iv) Group 5: 20- day-old flies starved for seven days after the last blood meal (starved old

flies). To investigate whether starvation affects the established midgut infection progress to

metacyclic infectious form in salivary glands, non-starved teneral (Group 3) and 20-day-old

(Group 6) flies at the time of the infective meal were deprived of blood meal after the midgut

establishment period (i.e. 10 days after the infective meal).

For T. b. gambiense, one series of experimental infection was conducted at the

“Laboratoire de Recherches et de Coordination sur les Trypanosomes (IRD-CIRAD)” in

Montpellier (France). Five experimental groups of G. palpalis gambiensis (for rearing

conditions see Ravel et al., 2006) were infected with the T. b. gambiense strain i.e. i) non-

starved teneral flies (newly emerged flies less than 32 hours old), ii) teneral flies that were

starved for four days, iii) 12-day-old flies starved for two days after the last blood meal (non-

starved adults), iv) 12- day-old flies starved for five days after the last blood meal (starved

adults) and v) 13-day-old flies starved for 10 days after the last blood meal. In a second series

of experimental infections, T. b. gambiense MBA, T. b. gambiense NHOM/INRB/2007 ) and

T. b. gambiense S12.9.5 were used to infect non-starved and starved teneral and adult flies G.

m. morsitans (the division into groups were similar as described above for T. b. rhodesiense).


35

2.2.3. Tsetse flies infective blood meal and maintenance

For each experimental group, flies were given a single infective blood meal on anesthetized

trypanosome-infected mice (strain NMRI for experiments conducted at ITM and Balb/cj for

experiments conducted in Montpellier) (one batch of 40 flies per mouse) showing a

parasitaemia between 106.9 and 108.1 trypanosomes/ml of blood. Mice were immune-

suppressed with cyclophosphamide (Endoxan, 300 mg/kg). The parasitaemia of each mouse

was checked using Herbert and Lumsden’s (1976) method. Mice were anesthetized by an

intraperitoneal injection of 0.1 mL / 10 g body weight of a xylazine/ketamine mixture (1mL

of Ketamine 10% + 0.4 mL of Rompun 2% + 8.6 mL of sterile water for injection). After the

infective blood meal, only engorged flies were retained and afterwards fed three times per

week on healthy rabbits until two days before dissection.

Flies of Groups 3 and 6 (T. b. rhodesiense infection) were fed 3 times per week on

clean rabbits for a period of 10 days (following infective blood meal) corresponding to the

trypanosome establishment period in the tsetse fly’s midgut (Van Den Abbeele et al., 1999).

After this 10-days period, they were deprived of blood feeding for seven consecutive days. At

the end of the seven days starvation period, the normal feeding regimen was resumed until

two days before dissection. In order to avoid re-infection of the flies during the maintenance

feeding, the rabbits were replaced at weekly intervals.

2.2.4. Tsetse flies dissection and infection rates

Thirty days after the trypanosome-infected meal, the midgut and salivary glands of all

surviving flies were dissected using the method described by Lloyd and Johnson (1924) and

examined by phase contrast microscopy at 400x magnification for the presence of

trypanosome infections. The overall infection rates were calculated as the Intrinsic Vectorial

Capacity (IVC) (Le Ray, 1989). It was calculated as defined in section 1.1 (Chapter 1).

Animal ethics approval for the experiment infections was obtained from the Ethics

Committee of the Institute of Tropical Medicine, Antwerp, Belgium (Refs: PAR003-MC-K-

Try and PAR004-MC-M-Try).

2.2.5. Statistical analysis

The effect of starvation on (i) procyclic midgut infection (p), (ii) maturation to metacyclic

infection in the salivary glands (m) and (iii) the overall metacyclic infection rate (IVC) in

teneral and older flies were analysed in three separate logistic regressions in STATA Version

9.2 (StataCorp, Inc., College Station, TX, USA). The response variables were the proportion


36

of infected flies whereas starvation, fly age class (older or teneral) were used as explanatory

variables. Cluster effect resulting from flies infected on the same mouse and maintained in the

same cage was taken into account.

2.3. Results

2.3.1. Infections with T. b. gambiense

A total of 127 teneral and 160 adult flies G. p. gambiensis were dissected 45 days after the

uptake of blood meal containing T b. gambiense S1/1/6. Of these, only three flies were found

positive in the midgut, with only few trypanosomes present. None of the dissected flies were

salivary glands-positive. Moreover, experimental infections conducted at ITM using the G. m

.morsitans and other T. b. gambiense strains, resulted in only 8 flies with a midgut infection

on a total of 327 dissected flies.

2.3.2. Infections with T. b. rhodesiense

A total of 974 flies (i.e. 424 flies that were fed an infectious blood meal in the teneral state

and 550 flies when 20-days-old) were dissected to determine their infection status at the

midgut and salivary glands levels. The proportion of flies with an established procyclic

midgut infection was high and was significantly increased when the flies were starved before

the infective blood meal (0.73 versus 0.55 and 0.17 versus 0.06 respectively in teneral and 20-

day-old flies) (Table 2.1). Over 50% and 30% of procyclic infections, respectively in teneral

and older flies, matured into metacyclic infectious trypanosomes in salivary glands. Starvation

did not affect significantly the maturation of the procyclic infection in teneral flies nor in

older flies (Table 2.2). As an overall result, the vectorial capacity of the nutritionally-stressed

tsetse flies for the T. b. rhodesiense parasite was increased, especially in the 20-day-old fly

group starved before the uptake of the parasite.


37

Table 2.1: Proportion (+95% confidence interval) of starved and non-starved teneral and 20-

day-old G. m. morsitans male flies that developed a trypanosome infection with T. b.

rhodesiense in the midgut (procyclic stage) and salivary glands (mature, metacyclic stage), 30

days after the infected blood meal. In Group 3 (teneral) and Group 6 (older), flies were

starved for seven consecutive days after the trypanosome midgut establishment period (10

days after the infected blood meal) whereas in Group 2 (teneral) and Group 5 (older) flies

were starved respectively for four and seven consecutive days before trypanosome-infected

blood meal.

Proportion of infected flies/ maturation

Fly groups

Number of flies

dissected Procyclics (p) Maturation (m) IVC*

Group 1 156 0.55 (0.45–0.65) 0.58 (0.44-0.71) 0.32

Group 2 130 0.73 (0.65–0.79) 0.53 (0.45-0.61) 0.39

Group 3 138 0.71 (0.64-0.76) 0.57 (0.46-0.67) 0.40

Group 4 212 0.06 (0.04-0.09) 0.35 (0.18-0.56) 0.02

Group 5 224 0.17 (0.13-0.21) 0.41 (0.31-0.51) 0.07

Group 6 114 0.07 (0.03-0.12) 0.37 (0.19-0.60) 0.03

* IVC = p x m = the number of dissected flies that developed a metacyclic infection

Table 2.2: Significance (P value) of the difference between the proportion of male G. m.

morsitans with a procyclic or metacyclic infection and IVC compared to non-starved flies

(Group 1 and Group 4 served as reference for teneral and older flies, respectively).

P values

Fly group Procyclic infection Maturation IVC

Group 2 0.012 0.559 0.369

Group 3 0.023 0.874 0.221

Group 5 < 0.001 0.651 0.002

Group 6 0.865 0.907 0.834

2.4. Discussion

Although T. b. gambiense S/1/1/6 has displayed immature and mature infections in a recent

experimental infection (Ravel et al., 2006), it failed in our experiments to infect either the

savannah or the riverine tsetse fly species. Other T. b. gambiense strains also failed to infect

the ITM G. m. morsitans flies. These results clearly demonstrate the difficulty in obtaining a


38

suitable T .b. gambiense-Glossina experimental model that allows the study of factors that

influence this parasite’s development. For the other human pathogenic species T. b.

rhodesiense our results showed that a recently isolated strain (T. b. rhodesiense Rumphi)

develops successfully in the tsetse fly G. m. morsitans. Moreover, the proportion of

metacyclic infected flies was substantially higher than those observed in similar studies.

Indeed, based on the outcome of infection experiments using the combination teneral G. m.

morsitans and T. b. rhodesiense, the proportion of metacyclic infected flies reported in the

literature was usually below 0.10 (Burtt, 1946; Harley, 1971; Gingrich et al., 1982a). In our

case, metacyclic infection rates in teneral flies were similar to the metacylic infection rates

obtained when infecting with T. b. brucei, a parasite that is considered to have a higher

transmissibility than T. b. rhodesiense (Welburn et al., 1995; Kubi et al., 2006). Importantly,

starvation occurring before the uptake of the parasite increased the overall metacyclic

infection rate, especially of the 20-day-old flies (up to 7% of metacyclic infected flies),

confirming the preliminary findings of Gingrich et al. (1982b). However, contrary to their

observations, the elevated infection rate was solely the result of a significantly lower barrier

for trypanosome establishment at the midgut level, and this was observed both in teneral as

well as in the 20-day-old flies group. However, we should be cautious to compare results

from similar trypanosome-tsetse fly infection experiments as a completely different

experimental set-up was used (i.e. differences at the level of the T. b. rhodesiense strain,

origin of G. morsitans flies, starvation periods and trypanosome infection procedures).

Moreover, our observations are based upon a solid number of experimental flies (130 to 224

flies in different groups) whereas the experiment of Gingrich was performed with only a

limited number of flies (18 to 65 flies in different groups). The enhancement of the T. b.

rhodesiense transmission by nutritionally stressed flies confirms previous observations made

with T. congolense and T. b. brucei using an identical experimental set-up (Kubi et al., 2006).

These results suggest that tsetse-related factors that constitute the developmental barrier in the

fly midgut are suppressed when the flies are under a nutritional stress. Since tsetse immune

system components are taught to be involved in their natural refractoriness (Lehane et al.,

2004), it is possible that starvation reduces the immunological responses of the fly to

eliminate the invading trypanosome.

Our results clearly indicate that the recently isolated T. b. rhodesiense RUMPHI strain is

highly transmissible. Besides the fact that G. m. morsitans flies are good vectors for different

trypanosomes (Kubi et al., 2006; Van Den Abbeele et al., 1999), the trypanosome strain itself


39

may also have intrinsic properties determining its high level of transmissibility. Indeed,

various field and experimental studies have revealed considerable differences in

transmissibility between different trypanosomes species and within a trypanosome species

between different strains. In T. congolense, for example, it has been demonstrated that highly

virulent and drug resistant strains have increased transmissibility by tsetse flies (Masumu et

al., 2006; Van den Bossche et al., 2006). Moreover, infectivity to man has been shown to

influence trypanosome transmissibility by tsetse flies with human serum-resistant T. b.

rhodesiense strains maturing far more less than human serum-sensitive T. b. brucei

trypanosomes (Welburn et al., 1995).

The epidemiological repercussions of these findings are substantial. Contrary to

previous observations, highly transmissible T. b. rhodesiense strains do occur. Although the

presence of such strains is difficult to detect in areas where trypanotolerant game animals

constitute the main reservoir of the parasite, they could play an important role in the spread of

the parasite in game and, perhaps more important, livestock populations. Such highly

transmissible T. b. rhodesiense strains when present in the livestock population could result in

high infection rates of tsetse flies in peri-domestic areas and cause epidemics in people. The

latter happened in Soroti District of eastern Uganda, where an outbreak of T. b. rhodesiense

sleeping sickness was attributed to cattle infected with this parasite (Fèvre et al. 2001).

Moreover, the observed increased susceptibility in starved flies may explain the focal

distribution character of sleeping sickness caused by T. b. rhodesiense. It is possible that in

areas where this parasite exist and where environmental conditions such as high ambient

temperature induce nutritional stress in tsetse, the number of flies becoming infected may

increase and subsequently induce the occurrence of disease epidemics or facilitate their

spread. Further studies are required to develop a T. b. gambiense transmission model to

conduct similar transmission experiments.


40

2.5. References

Barrett, M.P., 1999. The fall and rise of sleeping sickness. Lancet 353: 1113-1114


rhodesiense in Glossina morsitans. Annale of Tropical Medicine and Parasitology 40: 18-28

Elsen, P., Van Hees, J., De Lil, E., 1993. L'historique et les conditions d'élevage des lignées de glossines (Diptera, Glossinidae) maintenues à l'Institut de Médecine Tropical Prince Léopold d'Anvers. Journal of African Zoology 107: 439-449

Fairbairn, H., Watson, H.J., 1955. The transmission of Trypanosoma vivax by Glossina palpalis. Annal of Tropical Medicine and Parasitology 49: 250-259

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. Lancet 358: 625-628

Gingrich, J.B., Ward, R.A., Macken, L.M., Schoenbechler, 1982a. Trypanosoma brucei

rhodesiense (Trypanosomatidae): Factors influencing rates of a recent human isolate in the tsetse Glossina morsitans (Diptera: Glossinidae). Journal of Medical Entomology 19: 268-274

Gingrich, J.B., Ward, R.A., Macken, L.M., Esser, K.M., 1982b. African sleeping sickness: new evidence that mature tsetse flies (Glossina morsitans) can become potent vector. Transactions of the Royal Society of Tropical Medicine and Hygiene 76: 479-481

Harley, J.M.B., 1971. Comparison of the susceptibility to infection with Trypanosoma

rhodesiense of Glossina pallidipes, G. morsitans, G. fuscipes and G. brevipalpis. Annale of Tropical Medicine and Parasitology 65:185-189

Herbert, W.J., Lumsden, W.H.R., 1976. Trypanosoma brucei. A rapid matching method for estimating the host’s parasitemia. Experimental Parasitology 40: 427-431

Hirumi, H., Hirumi, K., 1989. Continuous cultivation of Trypanosoma brucei blood stream forms in a medium containing a low concentration of serum protein without feeder cell layers. The Journal of Parasitology 75: 985-989

Jamonneau, V., Ravel, S., Kiffi, M., Kaba, D., Zeze, D.G., Ndri, L., Sane, B., Coulibaly, B., Cuny, G., Solano, P., 2004. Mixed infections of trypanosomes in tsetse and pigs and their epidemiological significance in a sleeping sickness focus of Côte d’Ivoire. Parasitology 129: 693-702

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Van den Bossche, P. 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392


41




Llyod, L., Johnson, W.B., 1924. The trypanosome infections of tsetse flies in northern Nigeria and a new method of estimation. Bulletin of Entomological Research 14: 265-288

Masumu, J., Marcotty, T., Ndeledje, N., Kubi, C., Geerts, S., Vercruysse, J., Dorny, P., Van den Bossche, P., 2006. Comparison of the transmissibility of Trypanosoma congolense strains, isolated in a trypanosomiasis endemic area of eastern Zambia, by Glossina morsitans morsitans. Parasitology 133: 331-334

Ndegwa, P.N., Irungu, L.W., Moloo, S.K., 1992. Effect of puparia incubation temperature: increased infection rates of Trypanosoma congolense in Glossina morsitans centralis, G. fuscipes fuscipes and G. brevipalpis. Medical and Veterinary Entomology 6:127-130

Ravel, S., Patrel, D., Koffi, M., Jamonneau, V., Cuny, G., 2006. Cyclical transmission of Trypanosoma brucei gambiense in Glossina palpalis gambiensis displays great differences among field isolates. Acta Tropica 100: 151-155

Taylor, A.W., 1932. The development of West African strains of Trypanosoma gambiense in Glossina tachinoides under normal laboratory conditions and at raised temperatures. Parasitology 24: 401-418

Van Den Abbeele, J., Claes, Y., Van Bockstaele, D. and Le Ray, D., 1999. Trypanosoma brucei spp. Development in the tsetse fly: characterization of the post-mesocyclic stages in the foregut and proboscis. Parasitology 118, 469-478.

Van den Bossche, P., Akoda, K., Kubi, C., Marcotty, T., 2006. The transmissibility of Trypanosoma congolense seems to be associated with its level of resistance to isometamidium chloride. Veterinary Parasitology 135: 365-367

Welburn, S.C., Maudlin, I., Milligan, P.J.M., 1995. Trypanozoon: infectivity to human is linked to reduced transmissibility in tsetse. I. Comparison of human serum-resistant and human serum-sensitive field isolates. Experimental Parasitology 81: 404-408

Chapter 3

Maturation of a Trypanosoma brucei infection to the infectious

metacyclic stage is enhanced in nutritionally stressed tsetse flies

Akoda K., Van den Bossche P., Lyaruu E.A., De Deken R., Marcotty T., Coosemans M. and

Van Den Abbeele J. (2009). Maturation of a Trypanosoma brucei infection to the infectious

metacyclic stage is enhanced in nutritionally stressed tsetse flies. Journal of Medical

Entomology (In press)

Chapter 3: Nutritional stress and T.b.brucei midgut infection maturation

43

3.1. Introduction

The developmental cycle of African trypanosomes in tsetse flies (Diptera: Glossinidae) starts

when the insect feeds on a trypanosome-infected mammalian host. Then, the ingested

parasites undergo a series of developmental stages including the establishment of a procyclic

infection in the fly’s midgut as well as the upwards migration to the mouthparts (for T.

congolense) or salivary glands (for T. brucei spp.) where a complex differentiation takes place

to the final mature metacyclic stage. This end stage is again infectious for a mammalian host

and will be transmitted at every blood feeding event of the infected tsetse fly. The existence of

developmental barriers is evidenced by the low proportion of trypanosome infections in the

midgut of experimentally infected flies and the fact that only a limited proportion of these

midgut infections will finally give rise to a mature infection especially in the case of T. brucei

spp (Roditi and Lehane, 2008). The proportions of infected flies in a tsetse population as well

as the age-specific susceptibility are important factors that affect the epidemiology of tsetse-

transmitted trypanosomiasis. Tsetse flies are considered to be most susceptible for T.

congolense and T. brucei spp. infection in their teneral state i.e. before their first blood meal

(Welburn and Maudlin, 1992). The large majority of infected tsetse flies are considered to

have acquired the infection during their first blood meal, while those that do not become

infected at an early stage are not believed to contribute to the trypanosome infection rate of

the tsetse population. However, under specific physiological conditions, both teneral and

older flies can become more susceptible to develop a trypanosome infection. Indeed, Kubi et

al. (2006) showed that nutritional deprivation of tsetse flies lowers the developmental barrier

to establish a trypanosome infection, particularly at the tsetse midgut level. However, the

effect of this nutritional stress on the maturation of an established procyclic infection to the

infective metacyclic stage has not been examined thoroughly. Therefore, the experimental

work presented in this study focused on the effect of starvation of tsetse flies on the

maturation of a T. brucei procyclic midgut infection into a metacyclic salivary gland

infection.

3.2. Materials and Methods

3.2.1. Tsetse flies and trypanosome strain

Freshly emerged male Glossina morsitans morsitans Westwood flies (less than 32 h-old) were

used in this experiment. Their origin and rearing technique were described in chapter 2

section 2.2.2. Trypanosoma brucei brucei AntAR1 strain derived from the stock EATRO

1125 (Le Ray et al., 1977) was used in the experiments.


44

3.2.2. Experimental design

Twenty six cages of 40 newly emerged male flies each were given a single blood meal on

anesthetized trypanosome-infected mice (NMRI) (one cage per mouse) showing a

parasitaemia of 108.1 - 108.4 trypanosomes/mL of which more than 50% were short-stumpy

bloodstream forms. Mice were anesthetized by an intraperitoneal injection of a

xylazine/ketamine mixture. After the infected blood meal, unfed flies were removed. During

a period of 10 days corresponding to the trypanosome establishment period in the tsetse fly’s

midgut (Van Den Abbeele et al., 1999), all these flies were fed three times per week on

uninfected rabbits. After this 10-days period, flies were divided into two experimental groups,

each containing 13 replicates (i.e. cages of flies fed on a different infected mouse). Group 1

comprised starved flies, that were deprived of blood feeding for seven consecutive days

whereas Group 2 consisted of non-starved flies that continued a normal feeding regimen of

three times per week. At the end of the seven-day starvation period for Group 1, this normal

feeding regimen was resumed until two days before dissection. To avoid re-infection of the

flies during the maintenance feeding, the rabbits were replaced at weekly intervals. Thirty

days after the infective meal, surviving flies were dissected and midgut and salivary glands

were examined by phase-contrast microscopy (400x) to determine the presence of

trypanosomes. The proportion of midgut procyclic infections (immature infection) was

calculated as the proportion of dissected flies that developed a trypanosome infection in the

midgut. The maturation rate was calculated as the proportion of immature infections that

developed into mature infections in the salivary glands

Statistical analyses were carried out in STATA Version 9.2 (StataCorp, Inc., College

Station, TX, USA) using a logistic regression. The proportions of dissected flies showing

infection in the midgut and in the salivary glands were tested separately. Starvation was taken

as explanatory variable. Clusters resulting from flies infected on the same mouse and

maintained in the same cage were considered as primary sampling units. The maturation was

analyzed in a similar way but on a dataset restricted to midgut infected flies.

Animal ethics approval for the experimental infections was obtained from the Ethics

Commission of the Institute of Tropical Medicine, Antwerp, Belgium (Refs: PAR003-MC-K-

Try and PAR004-MC-M-Try).


45

3.3. Results and Discussion

Of a total of 1040 newly emerged male flies, 755 (72.5%) ingested a trypanosome-infected

blood meal and were retained for the experiment. Throughout the experiment, the mortality

rates were similar in both experimental groups, 8.8% and 5.5% respectively in Group 1

(starved) and Group 2 (non-starved) flies. At day 30, a total of 663 flies were dissected to

determine the infection status in the midgut and salivary glands (Table 3.1). In both groups,

the percentage of midgut-infected flies was high and was not significantly different (50% and

45% respectively, P = 0.2). However, in Group 1 flies a significantly higher number of these

midgut-infected flies developed a mature metacyclic infection in the salivary glands (P =

0.002). These results clearly demonstrate that an established T. brucei midgut infection

remains persistent even during a period in which the tsetse fly is exposed to a high nutritional

stress. In addition, our data suggest that the maturation of this persistent procyclic midgut

infection is significantly enhanced by the fly starvation resulting in a higher proportion of

flies that finally carry mature, infectious metacyclic trypanosomes. Previous studies have

already demonstrated that a period of starvation of newly emerged and older tsetse flies

before the infective blood meal increases the susceptibility of these flies to establish a T.

congolense or T. b. brucei midgut infection (Kubi et al., 2006). However this is, to our

knowledge, the first time to demonstrate unambiguously that starvation of tsetse flies also

significantly affects the further development of established T. b. brucei procyclic

trypanosomes to metacyclic forms. However, this contrasts with what was observed in the

case of T. b. rhodesiense where starvation occurring during the maturation process lowered

the establishment barrier but did not affect the progress of established infection into

metacyclic infectious forms (Chapter 2).

The underlying mechanism for trypanosome maturation enhancement for the T. brucei

brucei parasite is not clear. We could hypothesize that, as a result of the nutritional stress of

the tsetse fly, the presence of factors that prevent the parasite to mature is reduced or that the

level of factors that promote maturation is increased. However, the nature of these factors

affecting the maturation of a midgut T. brucei infection remains unknown. In G. morsitans

flies, maturation of T. brucei spp. is greatly affected by fly sex with male tsetse flies

producing significantly more mature trypanosome infections than female flies (Welburn and

Maudlin, 1999). In a recent study, Macleod et al. (2007) suggested that oxidative stress (i.e.

presence of reactive oxygen species (ROS) such as nitric oxide) might be involved in the

triggering of the maturation process of a T. b. brucei midgut infections in tsetse. In the same


46

study, the authors demonstrated that pregnancy in female tsetse flies has a detrimental effect

on the parasite maturation which they attributed to the altered physiology/biochemistry in the

pregnant females and to a reduction of free nutrients available for the trypanosomes.

According to our study, on the other hand, the trypanosome maturation process was enhanced

when male midgut-infected tsetse flies were deprived of nutrients as a result of starvation.

However, it is highly probable that the above described studies cannot be simply compared

and that two different mechanisms are at play. Indeed, besides the specific physiological and

biochemical status of pregnant females (Attardo et al., 2006), it can be assumed that the

continuous intrauterine nourishment of the larva may result in a decrease of specific nutrients

whereas the extensive starvation of the male flies will result in a general and substantial

decrease of all free nutrients. Other studies have shown that specific immune responses of the

tsetse fly against the trypanosome parasite interfere with the development of the trypanosome

in the fly (Boulanger et al., 2002, Aksoy et al., 2003, Lehane et al., 2004, Attardo et al.,

2006). Since nutritional stress affects the immune status of G. morsitans flies (Akoda et al.,

2009), this could also be a contributing factor to the enhanced maturation of the established

procyclic trypanosomes.

Table 3.1: Proportion (+95% confidence interval) of G. m. morsitans male flies that

developed a trypanosome infection with T.brucei brucei AnTAR 1 in the midgut (procyclic

stage) and salivary glands (mature, metacyclic stage), 30 days after the infected blood meal.

In Group 1, flies were starved for seven consecutives days after the trypanosome midgut

establishment period (10 days after the infected blood meal) whereas Group 2 flies continued

to be fed three times a week

Proportion of infected flies

Groups

Total number

flies dissected Midgut (p) Maturationa (m) IVCb

Group 1 332 0.50 (0.44-0.55) 0.63c (0.55-0.69) 0.32c (0.26-0.36)

Group 2 331 0.45 (0.39-0.50) 0.47 (0.38-0.54) 0.21 (0.16-0.25)

a: Maturation = proportion of midgut infected flies that developed a mature, metacyclic infection in

the salivary glands

b: IVC = p x m = the number of dissected flies that developed a metacyclic infection

c: p=0.002


47

The findings of this study, together with those from Kubi et al. (2006), contribute to a better

understanding of the dynamics of T. brucei transmission in a natural tsetse population.

Indeed, it seems that a range of environmental factors that cause nutritional stress not only

make young and old tsetse flies more susceptible to establish a trypanosome midgut infection

(Kubi et al., 2006) but also boost the maturation of a midgut infection to the infectious stage

that would not have matured under normal circumstances. Since the T. brucei spp. infection

rates in natural tsetse fly populations are usually very low (Hide, 1999; Waiswa et al, 2006),

any significant increase in the overall mature infection rate may result in the enhanced

transmission of this parasite within a susceptible host population. As such, factors enhancing

trypanosome development may contribute to the maintenance and/or activation of a sleeping

sickness focus in an endemic area.

In the view of tsetse females’ higher longevity and their ensuing role in trypanosome

transmission (Welburn and Maudlin, 1999), it would be interesting to determine whether the

ability to transmit trypanosomes of female G. morsitans flies is similarly affected by

starvation. Moreover, similar experimental studies using other tsetse fly – trypanosome

transmission models are required to broaden our appreciation of the impact of nutritional

stress on the vector competence of other tsetse fly species.


48

3.4. References

Akoda, K., P. Van den Bossche, T. Marcotty, C. Kubi, R. De Deken,, J. Van Den Abbeele. 2009. Nutritional stress affects the tsetse fly’s immune gene expression. Medical and

Veterinary Entomology 23:195–201

Aksoy, S., W.C. Gibson,, M.J. Lehane. 2003. Interactions between tsetse trypanosomes with implications for the control of trypanosomiasis. Advance in Parasitology 53:1-83.

Attardo,G.M., Lohs,C., Heddi,A., Alam, U.H., Yildirim,S., Aksoy, S., 2008. Analysis of milk gland structure and function in Glossina morsitans: Milk protein production, symbiont populations and fecundity. Journal of Insect Physiology 54: 1236-1242.

Attardo, G.M., P. Strickler-Dinglasan, S.A.H., Perkin, E. Caler, M.F. Bonaldo, M.B. Soares, N. El-Sayeed, Aksoy, S., 2006. Analysis of fat body transcriptome from the adult tsetse fly, Glossina morsitans morsitans. Insect Molecular and Biology 15: 411-424.

Boulanger, N., Brun, R., Ehret-Sabatier, L., Kunz, C., Bulet, P., 2002. Immunopeptides in the defense reactions of Glossina morsitans to bacterial and Trypanosoma brucei brucei infections. Insect Biochemistry and Molecular Biology 32: 369-375.

Hide,G., 1999. History of Sleeping Sickness in East Africa. Clinical Microbiology Review 12: 112-125.

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Van den Bossche, P., 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392.

Lehane, M.J., Aksoy, S., Levashina, E., 2004. Immune responses and parasite transmission in blood-feeding insects. Trends Parasitology 20:433-439.

Le Ray, D., Barry, J.D., Easton, C., Vickerman, K., 1977. First tsetse fly transmission of the ‘Antat’ serodeme of Trypanosoma brucei. Annales de la Société Belge de Médécine

Tropicale 57: 369-381.

Macleod, E.T., Darby, A.C., Welburn, S.C., 2007. Factors affecting trypanosome maturation in tsetse flies. PLOS One, 2, e239. doi:10.137/journal.pone.0000239.

Roditi, I., Lehane, M.J., 2008. Interactions between trypanosomes and tsetse flies. Current

Opinion in Microbiology 11:345-351.

Van Den Abbeele, J., Claes, Y., Van Bockstaele, D., Le Ray, D., Coosemans, M., 1999. Trypanosoma brucei spp. Development in the tsetse fly: characterization of the post-mesocyclic stages in the foregut and proboscis. Parasitology 118: 469-478.

Waiswa,C., Picozzi, K., Katunguka-Rwakishaya, E., Olaho-Mukani,W. Musoke,R.A., Welburn, S.C., 2006. Glossina fuscipes fuscipes in the trypanosomiasis endemic areas of south eastern Uganda: Apparent density, trypanosome infection rates and host feeding preferences. Acta Tropica 99: 23-29.


49

Welburn, S.C., Maudlin, I., 1992. The Nature of the teneral state in Glossina and its role in the acquisition of trypanosome infection in tsetse. Annal of Tropical Medicine and

Parasitology 86: 529-536.

Welburn, S.C., Maudlin, I., 1999. Tsetse-trypanosome interactions: Rites of passage. Parasitology Today 15: 399-403

Chapter 4

Nutritional stress of adult female tsetse flies (Diptera: Glossinidae)

affects the susceptibility of their offspring to trypanosomal infections

Akoda K., Van Den Abbeele J. , Marcotty T. , De Deken R., Sidibé I. and Van den Bossche P.

(2009). Nutritional stress of adult female tsetse flies (Diptera: Glossinidae) affects the

susceptibility of their offspring to trypanosomal infections. Acta Tropica 111: 263–267

Chapter 4: Adult female tsetse starvation & their offspring’s susceptibility

51

4.1. Introduction

In Africa, tsetse-transmitted trypanosomiasis poses a serious threat to animals and humans.

About 10 million km2 of sub-Saharan Africa, extending over 37 countries, are infected by

tsetse flies. The epidemiology of this important disease is determined largely by the

proportion of infected tsetse flies. A range of intrinsic and extrinsic factors among which

environmental conditions, have been identified to affect the tsetse’s susceptibility to

trypanosomal infections and, thus affect the overall infection rate of the tsetse population

(Leak, 1999; Aksoy et al., 2003 ; Macleod et al., 2007). For example, infection rates in tsetse

flies are positively correlated with ambient temperature to which puparia or adult flies are

exposed (Taylor, 1932; Burtt, 1946; Fairbairn and Watson, 1955; Ndegwa et al., 1992).

Moreover, high ambient temperatures shorten the duration of the development cycle within

tsetse (Fairbairn and Culwick, 1950). The underlying mechanisms for the observed effect of

ambient temperature on the susceptibility to infection are not well understood. The nutritional

status of the tsetse fly at the time of the infective bloodmeal also affects its susceptibility to

infections with Trypanosoma congolense or Trypanosoma brucei brucei. Indeed, a period of

starvation (4 days for teneral flies and 7 days for adult flies) lowers the developmental barrier

for a trypanosome infection, especially at the midgut level of the fly (Kubi et al., 2006). Here,

it has been hypothesized that the reduction in the fat body reserve as a result of starvation

reduces the fly’s ability to mount a competent immune response against the invading

trypanosomes. It has been suggested that the tsetse’s innate immune response affects the

establishment and maturation of trypanosomes in the vector (Hao et al., 2001; Boulanger et

al., 2002; Lehane et al., 2004; Attardo et al. 2006). This immune response is determined by a

range of antimicrobial peptides among which attacin, defensin, cecropin and diptericin that

are mainly synthesized in the fat body in response to a ‘foreign’ micro-organism. Hence, all

factors that reduce the fat body level of a teneral or adult fly may result in concomitant

changes in the fly’s vector competence. In this paper, we investigated the effect of nutritional

stress of reproducing tsetse females on the susceptibility of their offspring to trypanosomal

infection. Results of this study may explain how changes in the tsetse’s environment, that

cause considerable stress, may have repercussions on the epidemiology of tsetse-transmitted

human or animal trypanosomiasis.


52

4.2. Materials and methods

4.2.1. Tsetse flies

Tsetse flies Glossina morsitans morsitans Westwood from the colony maintained at the

Institute of Tropical Medicine (Antwerp, Belgium) were used in the experiments. The origin

of this tsetse colony and rearing conditions were described by Elsen et al. (1993). Flies from

this colony were fed 4 times a week on healthy rabbits (= non-starved colony). In addition to

the main colony, a second colony of 2220 adult reproducing female flies was established and

nutritionally stressed by feeding the flies only once a week on healthy rabbits (= starved

colony) for 9 weeks. Pupae produced by the female flies of both colonies were collected daily

and weighed.

4.2.2. Trypanosome infection of tsetse flies

The clonal strains of Trypanosoma congolense IL 1180 (Geigy and Kauffmann, 1973) and

Trypanosoma brucei brucei AntAR1 (Le Ray et al., 1977), were used to infect the

experimental flies. Series of 30 male flies (less than 32h old) that emerged from pupae

produced by females of the starved or non-starved colony were given a single bloodmeal on

trypanosome infected anesthetized mice (NMRI). The mice were infected with either T.

congolense or T. b. brucei showing a parasitaemia of 106.9 or 108.1 trypanosomes per mL

respectively. Mice were anesthetized with a mixture of Anesketin® and Rompun®. Only fully

engorged flies were retained and fed afterwards on healthy rabbits three times per week. To

avoid re-infection of the flies, these rabbits were replaced at weekly intervals.

4.2.3. Tsetse infection rate

Twenty-one days (for infections with T. congolense) or 30 days (for infections with T. b.

brucei) after the infective bloodmeal, all surviving flies were dissected using the method

described by Lloyd and Johnson (1924). Microscopical examination of the midgut and

mouthparts or midgut and salivary glands were conducted to determine the presence of T.

congolense or T. b. brucei respectively.

The proportion of immature (Midgut) infections was calculated as the proportion of dissected

flies that had a trypanosomal infection in the midgut whereas the maturation rate was

calculated as the proportion of midgut infection that developed into a mature infection in the

mouthparts (for T. congolense) or salivary glands (for T. b. brucei).


53

4.2.4. Determination of the fat content in the offspring

Thirty (30) male flies that emerged from pupae of the starved or non-starved colony were

killed immediately after emergence, their legs and wings were removed and the carcasses

were dried at 80 °C to constant weight. Afterwards the dry weight of pools of three flies was

determined. Fat body was extracted through chloroform extraction of lipids for three days

with daily changes of chloroform (Langley et al., 1990). After fat extraction and drying to

constant weight, the residual dry weight of the pools of three flies was determined. The fat

content of the flies was calculated as the difference between the dry and residual dry weights.

Weighing was done using a Sartorius® analytical balance (Sartorius AG, Göttingen, Germany

(+/- 0.1 mg).

4.2.5. Immune peptide gene expression levels in the freshly emerged flies

To compare the baseline immune peptide gene expression level in male flies derived from

pupae of the starved or non-starved adult females, total RNA extraction followed by cDNA

synthesis and Quantitative real-time PCR were performed similarly as described in chapter 5.

4.2.6. Statistical data analysis

All statistical analyses were carried out in STATA Version 9.2 (StataCorp, Inc., College

Station, TX, USA). A linear regression was used to analyse the loss of weight in individual

flies. The difference in weight was used as the response and fly groups (flies emerging from

pupae produced by females of the starved and non-starved colony) as discrete explanatory

variables. The distribution of the residuals and the heteroskedasticity of the variance were

verified (P > 0.1). The proportions of infected flies in different experimental series were

compared using a robust logistic regression. Cluster effects resulting from flies infected by

feeding on the same mouse and maintained in the same cage were taken into account in the

model.

The effect of adult female starvation on the expression levels of attacin, defensin and cecropin

in emerging teneral male flies were analysed as performed in chapter 5. For each statistical

analysis, differences were considered significant when P < 0.05.

4.3. Results

During the study period, the starved colony produced 0.07 to 0.15 pupae per female per week

whereas the non-starved colony produced about 1.0 pupae per female per week. Many

abortions were observed in the starved colony. The pupal mean weight ranged between 20 and

22 mg in the starved colony while the pupal mean weight in the non-starved colony was about


54

26 mg. The pupal emergence rate was 89% in the starved colony and 97% in the non-starved

female colony.

4.3.1. Nutritional stress and expression of immune peptide genes of the emerging flies

Male flies emerging from pupae produced by the non-starved females had an almost 2-fold

higher fat body content than teneral males emerging from pupae produced by the starved

females (2.5 ± 0.1 mg compared to 1.4 ± 0.4 mg, p<0.0001). The normalized gene expression

levels of the immune peptides attacin, defensin and cecropin were respectively 1.5, 1.8 and

2.0 times higher in teneral male flies from non-starved adult females (Figure 4.1). None of

these differences are, however, statistically significant.

4.3.2. Trypanosome infection rates

The proportion of teneral males feeding on the infected mice was 74.8% and 65.6% for

tenerals emerging from pupae produced by the starved and the non-starved females

respectively. Mortality rates of flies after the infective feed were 7.1% and 1.5% for males

emerging from pupae produced by starved and non-starved females respectively.

0

0,1

0,2

0,3

0,4

0,5

0,6

Non-starved adult

females

Starved adult females

No

rma

lize

d e

xp

ressio

n

Attacin

Defensin

Cecropin

Figure 4.1: Normalized expression levels of attacin, defensin and cecropin in male teneral G.

m. morsitans emerging from non-starved or starved adult female flies and their 95%

confidence intervals (n = 24 and 20 abdomens respectively in non-starved and starved flies

group). Expression level of each immune gene was normalized against the value of the

housekeeping genes actin and tubulin.


55

The proportion of flies from the starved colony that developed a mature infection with T.

congolense (Table 4.1) or T. b. brucei (Table 4.2) was significantly higher compared to flies

from the non-starved colony (p < 0.001). In T. congolense-infected flies, the proportion of

male flies that developed a midgut infection was significantly higher in the flies emerging

from the starved colony compared to flies from the non-starved colony (0.44 versus 0.27, p=

0.0001). The maturation rate of the midgut infections was about 100% in both experimental

groups. For infections with T. b. brucei, the proportion of flies that developed a midgut

infection did not differ significantly between the experimental groups (p = 0.067) but

maturation rate was significantly higher in males emerged from pupae produced by females of

the starved colony (0.55 versus 0.35) (Table 4.2).

Table 4.1: Proportion of male teneral G. m. morsitans emerging from starved or non-starved

females and infected with T. congolense IL1180

Proportion of flies infected in Experimental group

Batch of

flies

Number

dissected Midgut (p) Maturation (m)a IVCb

1 28 0.50 (14/28) 1.00 (28/28) 0.50

2 17 0.41 (7/17) 1.00 (7/7) 0.41

3 25 0.40 (10/25) 1.00 (10/10) 0.40

4 27 0.41 (11/27) 1.00 (11/11) 0.41

5 25 0.48 (12/25) 0.92 (11/12) 0.44

Male teneral flies

emerging from starved

adult females 6 17 0.41 (7/17) 1.00 (7/7) 0.41

1' 15 0.20 (3/15) 1.00 (3/3) 0.20

2' 22 0.32 (7/22) 1.00 (7/7) 0.32

3' 40 0.30 (12/40) 1.00 (12/12) 0.30

4' 46 0.26 (12/46) 1.00 (12/12) 0.26

5' 28 0.25 (7/28) 1.00 (7/7) 0.25

Male teneral flies

emerging from non-

starved adult females 6' 31 0.29 (9/31) 1.00 (9/9) 0.29

a: Maturation = proportion of midgut infected flies that developed a mature, metacyclic infection in the

mouthparts



56

Table 4.2: Proportion of male teneral G. m. morsitans emerging from starved or non-starved

females and infected with T. b. brucei AnTAR 1

Proportion of flies infected in

Experimental group

Batch of

flies

Number

dissected Midgut (p) Maturation (m)a IVCb

1 16 0.56 (9/16) 0.44 (4/9) 0.25

2 20 0.65 (13/20) 0.54 (7/13) 0.35

3 18 0.61 (11/18) 0.64 (7/11) 0.39

4 15 0.53 (8/15) 0.63 (5/8) 0.33

Male teneral flies

emerging from

starved adult

females 5 28 0.64 (18/28) 0.50 (9/18) 0.33

1' 20 0.55 (11/20) 0.18 (2/11) 0.10

2' 26 0.58 (15/26) 0.27 (4/15) 0.15

3' 25 0.40 (10/25) 0.60 (6/10) 0.24

4' 27 0.52 (14/27) 0.43 (6/14) 0.22

Male teneral flies

emerging from non-

starved adult

females 5' 43 0.58 (25/43) 0.28 (7/25) 0.16


salivary glands


4.4. Discussion

The starvation induced in the adult female tsetse flies substantially increased their mortality

rate and reduced their reproductive performance. Moreover, the mortality rate of the offspring

of those starved females was substantially higher compared to that of offspring of the non-

starved females. This suggests a reduced viability of male tsetse flies produced by female flies

submitted to this level of nutritional stress. This is not surprising considering the tsetse’s

adenotrophic viviparity and the importance of the uterine milk produced by the mother fly for

the development of the three larval stages in its uterus (Denlinger and Ma, 1974).

Consequently, the condition of the mother fly has direct repercussions for its offspring.

Environmental conditions vary seasonally and have a significant effect on the fat

content and size of adult tsetse flies (Glasgow and Bursell, 1960; Van den Bossche and

Hargrove, 1999) as well as on the body size and survival of their offspring (Jackson, 1952;

Phelps and Clarke, 1974; Dransfield et al., 1989). Based on the performance parameters of

the starved colony, the nutritional stress induced in this colony seems to mimic the seasonal


57

stress experienced by field populations of tsetse flies. Especially the reduced fat level of the

emerging teneral male flies may be attributed directly to the nutritional stress the female flies

were subjected to. In accordance with observations made by Kubi et al. (2006), the mature

trypanosomal infection rate of teneral males with low fat levels (emerging from pupae

produced by females from the starved colony) was significantly higher than the infection rates

of males with higher fat levels (emerging from pupae produced by the non-starved colony).

This increased susceptibility to trypanosomal infections could be attributed partially to a

reduced baseline immune status of the offspring as a result of the nutritional stress

experienced by the parent female fly. Such hypothesis is supported by Koella and Sörensen’s

(2002) observations that sufficient and good quality food is essential for good immune

defence in insects. Similarly, Siva-Jothy and Thompson (2002) concluded that a reduction in

resource availability can decrease the immune function of invertebrates. In our study, the

impact of starvation on the immune system was only examined for immune peptides that are

mainly produced systemically by the fat body and used by the tsetse’s defence mechanisms

against trypanosome infection (Hao et al., 2001; Boulanger et al., 2002). The fact that the

expression of these immune peptides genes was reduced, but not significantly so, could be

due to the way in which this expression was measured. Indeed, measurement of the

expression in the whole abdomen of the fly may mimic significant reductions in the

expression of immune peptides in organs where the trypanosomes develop such as the midgut.

Other factors that are induced by starvation and that may affect the development of a

trypanosomal infection in tsetse can, however, not be excluded. They include midgut lectins,

anti-oxidants and symbiotic associations in tsetse (Abubakar et al., 2006; Munks et al., 2005;

Macleod et al., 2007; Roditi and Lehane, 2008).

Although these observations are based entirely on laboratory experiments, the

outcome may contribute to a better understanding of the seasonal dynamics of trypanosomal

infection rates in field populations of tsetse flies. Indeed, the monthly metacyclic infection

rate of tsetse populations does undergo substantial variations over the year. Such variations

have been attributed to changes in the survival rate of the tsetse population or the availability

of infected hosts (Jordan, 1965; Harley, 1967). Our results suggest that seasonal stress

situations, resulting in changes in the susceptibility to trypanosomal infections of emerging

teneral flies, could also play a major role. Especially the hot dry season when ambient

temperatures are high constitutes a stressful period for tsetse flies. This is reflected in the

decrease in the fly’s body size and fat body level during this season (Jackson, 1952; Glasgow


58

and Bursell, 1960; Dransfield et al., 1989). The association between high ambient

temperatures, stress in tsetse flies and the concomitant increase in the tsetse susceptibility to

trypanosomal infections may offer an explanation for the often observed relationship between

high temperatures and high trypanosomal infection rates in tsetse. For example, Kinghorn and

Yorke (1912) and Burtt (1946) reported higher susceptibility to infection with T. b.

rhodesiense of flies emerging from pupae incubated at a high temperature. The same

association may explain partly Fairbairn’s (1948) observation that in the western G. morsitans

belt of Tanzania the monthly mean maximum temperature and the number of diagnosed cases

of sleeping sickness was significantly correlated. Finally and on a continental scale, a review

of surveys determining the infection rates of savannah species of tsetse flies conducted over a

period of 45 years also showed a positive correlation between infection rate of tsetse flies and

the mean annual temperature (Ford and Leggate, 1961). Nevertheless, further research is

required to determine the field implications of our laboratory observations.


59

4.5. References

Abubakar, L.U., Bulimo,W.D., Mulaa,F.J., Osir,E.O., 2006. Molecular characterization of a tsetse fly midgut proteolytic lectin that mediates differentiation of African trypanosomes. Insect Biochemistry and Molecular Biology 36: 344-352

Aksoy, S., Gibson, W., Lehane, M.J., 2003. Interactions between tsetse and trypanosomes with implications for the control of trypanosomiasis. Advance in Parasitology 53: 1-83

Attardo, G.M., Strickler-Dinglasan, P., Perkin, S.A.H., Caler, E., Bonaldo, M.F., Soares, M.B., El-Sayeed, N., Aksoy, S., 2006. Analysis of fat body transcriptome from the adult tsetse fly, Glossina morsitans morsitans. Insect Molecular Biology 15: 411-424.



rhodesiense in Glossina morsitans. Annal of Tropical Medicine and Parasitology 40: 18-28.

Denlinger, D.L., Ma, W.C., 1974. Dynamics of the pregnancy cycle in the tsetse fly Glossina

morsitans. Journal of Insect Physiology 20: 1015-1026.

Dransfield, R. D., Brightwell, R., Kiilu J., Chaudhury, M. F. B., Adabie, D. A., 1989. Size and mortality rates of Glossina pallidipes in the semi-arid zone of southwestern Kenya. Medical and Veterinary Entomology 3: 83-95.

Elsen, P., Van Hees, J., De Lil, E., 1993. L'historique et les conditions d'élevage des lignées de glossines (Diptera, Glossinidae) maintenues à l'Institut de Médecine Tropical Prince Léopold d'Anvers. Jouranal of African Zoology 107: 439-449.

Fairbairn, H., 1948. Sleeping sickness in Tanganyika territory, 1922-1946. Tropical Disease

Bulletin 45: 1-17.

Fairbairn, H., Culwick, A.T., 1950. The transmission of polymorphic trypanosomes. Acta

Tropica 7: 19-47.

Fairbairn, H., Watson, H.J., 1955. The transmission of Trypanosoma vivax by Glossina palpalis. Annal of Tropical Medicine and Parasitology 49: 250-259.


Society for Tropical Medicine and Hygiene 55: 383-397.

Geigy, R., Kauffman, M., 1973. Sleeping sickness survey in the Serengeti area (Tanzania) 1971. I. Examination of large mammals for trypanosomes. Acta Tropica 30: 12-23.

Glasgow, J. P., Bursell, E., 1960. Seasonal variations in the fat content and size of Glossina

swynnertoni Austen. Bulletin of Entomological Research 51: 705-713.


60


Sciences of the USA 98, 12648-12653.

Harley, J. M. B., 1967. Further studies on age and infection rate of Glossina pallidipes, G. palpalis fuscipes and G. brevipalpis in Uganda. Bulletin of Entomological Research 57: 459-477.

Jackson, C.H.N., 1952. Seasonal variations in the mean size of tsetse flies. Bulletin of


Jordan, A.M., 1965. The hosts of Glossina as the main factor affecting Trypanosome infection rates of tsetse flies in Nigeria. Transactions of the Royal Society for Tropical Medicine

and Hygiene 59: 423-431.

Kinghorn, A., York, W., 1912. On the influence of meteological conditions on the development of Trypanosoma rhodesiense in Glossina morsitans. Annal of Tropical


Koella, J.C., Sörensen, F.L., 2002. Effect of adult nutrition on the melanisation immune response of the malaria vector Anopheles stephensi. Medical and Veterinary

Entomology 16: 316-320

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Van den Bossche P., 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392.

Langley, P.A., Hargrove, J.W., Wall, R.L., 1990. Maturation of the tsetse fly Glossina

pallidipes (Diptera: Glossinidae) in relation to trap-orientated behaviour. Physiological

Entomology 15: 179-186.

Leak, S.G.A., 1999. Tsetse biology and ecology: their role in the epidemiology and control of trypanosomosis. Wallingord Oxon: CABI Publishing. 592 pp.

Lehane, M.J., Aksoy, S., Levashina, E., 2004. Immune responses and parasite transmission in blood-feeding insects. Trends Parasitology 20: 433-439.

Le Ray, D., Barry, J.D. Easton, C., Vickerman, K., 1977. First tsetse fly transmission of the ‘Antat’ serodeme of Trypanosoma brucei. Annale de la Société Belge de Médecine

Tropicale 57: 369-381

Llyod, L., Johnson, W.B., 1924. The trypanosome infections of tsetse flies in northern Nigeria and a new method of estimation. Bulletin of Entomological Research 14: 265-288.

Macleod, E.T., Maudlin, I., Darby, A.C., Welburn, S.C., 2007. Antioxidants promote establishment of trypanosome infections in tsetse. Parasitology 134: 827-831.

Munks, R.J., Sant’Anna, M.R., Grail, W., Gibson, W., Igglesden, T., Yoshiyama, M., Lehane, S.M., Lehane, M.J., 2005. Antioxidant gene expression in the blood-feeding fly Glossina morsitans morsitans. Insect Molecular Biology 14: 483-491.


61

Ndegwa, P.N., Irungu, L.W., Moloo, S.K., 1992. Effect of puparia incubation temperature: increased infection rates of Trypanosoma congolense in Glossina morsitans centralis, G. fuscipes fuscipes and G. brevipalpis. Medical and Veterinary Entomology 6: 127-130.

Phelps, R.J., Clarke, G.P.Y., 1974. Seasonal elimination of some size classes in male Glossina morsitans morsitans Westw. Bulletin of Entomological Research 64: 313-324

Roditi, I., Lehane, M.J., 2008. Interactions between trypanosomes and tsetse flies. Current

Opining Microbiology 11: 345-351.

Siva-Jothy, M., Thompson, J.J.W., 2002. Short-term nutrient deprivation affects immune function. Physiological Entomology 27: 206-212.

Taylor, A. W., 1932. The development of West African strains of Trypanosoma gambiense in Glossina tachinoides under normal laboratory conditions and at raised temperatures. Parasitology 24: 401-418.

Van den Bossche, P., Hargrove, J.W., 1999. Seasonal variation in nutritional levels of male tsetse flies Glossina morsitans morsitans Westwood (Diptera: Glossinidae) caught using fly-rounds and electric screens. Bulletin of Entomological Research 89: 381-387.

Chapter 5

Nutritional stress affects the tsetse fly’s immune gene expression

Akoda K., Van den Bossche P., Marcotty T., Kubi C., Coosemans M., De Deken R. and Van

Den Abbeele J. (2009). Nutritional stress affects the tsetse fly’s immune gene expression.

Medical and Veterinary Entomology 23, 195–201

Chapter 5: Nutritional stress & immune gene expression in tsetse fly

63

5.1. Introduction

Tsetse flies (Diptera: Glossinidae) are obligate blood feeding insects that cyclically transmit

African trypanosomes, protozoan flagellated parasites that cause nagana in livestock and

sleeping sickness in humans. The developmental cycle of the Trypanosoma congolense or T.

brucei parasite in the insect vector starts when the tsetse fly feeds on a trypanosome-infected

mammalian host. From this point on, the ingested parasite must undergo a series of

developmental stages that are located in the alimentary tract and the mouthparts or the

salivary glands of the fly. This complex journey within the tsetse fly infers two main barriers

which the parasite must overcome: i) the establishment of a procyclic infection in the tsetse

midgut within 3-5 days following the infective bloodmeal, and ii) migration and

differentiation to the final metacyclic infectious forms in the mouthparts or salivary glands,

depending on trypanosome species (Leak, 1999; Van Den Abbeele et al., 1999). These

developmental barriers are evidenced by the low proportion of trypanosome infections in the

midgut of experimentally infected flies and the fact that only a limited proportion of these

midgut infections will finally give rise to a mature infection. This implies that only a small

proportion of flies (both males and females) can eventually transmit the disease, a

phenomenon referred to as “refractoriness”. Our understanding of the tsetse-trypanosome

molecular interactions underlying this refractoriness is still limited. Many factors, including

midgut lectins, antioxidants and symbiotic associations in the tsetse fly, have been suggested

to affect the success or failure of the parasite’s development (Aksoy et al., 2003; Munks et al.,

2005; Abubakar et al., 2006; Macleod et al., 2007). In addition, the insect immune system has

been shown to play an important role in determining the fate of a trypanosome infection (Hao

et al., 2001). Indeed, tsetse flies are able to synthesize a range of antimicrobial peptides

(AMPs), such as attacin, defensin, cecropin and diptericin, in response to the presence of a

“foreign” micro-organism. This tsetse innate immune response is suggested to affect the

establishment and maturation of trypanosomes in the vector (Hao et al., 2001; Boulanger et

al., 2002; Lehane et al., 2004; Attardo et al., 2006; Hu and Aksoy, 2006). Moreover, Lehane

et al. (2008) provided evidence that an immune-responsive transferrin is also involved in the

tsetse-trypanosome interaction.

The proportions of infected flies in a tsetse population, as well as the age-specific

patterns susceptibility, are important factors that affect the epidemiology of trypanosomiasis.

In principle, the large majority of infected tsetse flies are considered to have acquired the

infection at their first bloodmeal as a teneral fly (newly hatched unfed fly). Older flies are


64

reported to be refractory to trypanosome infection and to contribute little to the overall

infection rate of a tsetse population. However, under specific physiological conditions, both

teneral and older flies can become more susceptible to trypanosome infection. Indeed,

starving teneral and 20-day-old tsetse flies for several days resulted in significant increases in

both T. congolense and T. brucei spp infections (Gingrich et al., 1982; Kubi et al., 2006). The

underlying mechanism that causes this important change in susceptibility has not yet been

clarified. Therefore, in this study, we evaluated whether starvation of the tsetse fly affects

uninduced and pathogen-induced innate immune responses in a way that may contribute to the

increased susceptibility of starved tsetse flies to trypanosome infection. As a limited

experimental read-out for the fly fat body-regulated immune response, the gene expression

levels of the AMPs attacin, defensin and cecropin were measured. These are AMPs regulated

by the immune deficiency (IMD) signalling pathway that are suggested to interfere with

trypanosome development in the tsetse fly (Hu and Aksoy, 2006).


5.2.1. Tsetse flies

Male Glossina morsitans morsitans Westwood tsetse flies from the colony maintained at the

Institute of Tropical Medicine (Antwerp, Belgium) were used throughout the experiments.

Their origin and rearing conditions were described in chapter 2.

Only male flies were used in the experimental set-up because an accurate fat analysis

of female tsetse flies is hampered by the presence of a developing larva in the female

abdomen as a result of the tsetse’s viviparous reproduction mode. Moreover, the molecular

interactions that play a role in the trypanosome development in the tsetse fly are assumed to

be similar in male and female flies; thus male flies can be used as an experimental model to

study these interactions and the factors affecting them.

5.2.2. Nutritional stress by starvation

In all experiments, four groups of flies were compared. The groups differed in age and

nutritional status. Group 1 comprised non- starved teneral flies (= newly emerged unfed flies)

aged < 32 h (TF0). Group 2 included teneral flies that had been starved for four days (TF4).

Group 3 consisted of 20-day-old flies starved for two days after their last bloodmeal (AD2).

Group 4 represented 20- day-old flies starved for seven days after their last bloodmeal (AD7).


65

5.2.3. Fat content determination

A total of 96 flies (24 flies per group) were used in this set of experiments. Flies were killed at

-20°C. Legs and wings were removed and carcasses were dried at 80°C to constant weight. To

increase the precision of the measurement of dry weights, flies were pooled by three for

weighing. Lipids were extracted using chloroform for three days (Langley et al., 1990).

Chloroform was replaced daily. After fat extraction and drying to constant weight, the

residual dry weight of the flies was determined. The fat content of the flies was calculated as

the difference between the dry and residual weights. All weightings were performed on a

Sartorius® analytical balance (Sartorius AG, Göttingen, Germany) (+ 0.1 mg).

5.2.4. Bacterial and trypanosomal challenge of tsetse flies

The four experimental groups of flies were immune-challenged by injecting bacteria or by

feeding the flies with trypanosome-infected blood.

To evaluate the bacteria-induced immune response in each of the experimental groups,

batches of 40 flies for each group were micro-injected either with 2 µL phosphate buffered

saline (PBS) (control) or with 2 µL of a suspension of live Escherichia coli (DH10B strain)

(OD620: approximately 0.6). Four days after the bacterial injection, whole abdomens were

removed and pooled by two for total RNA extraction.

For the trypanosomal challenge, batches of 40 flies for each group were given a

bloodmeal on anaesthetized mice (strain NMRI) showing a parasitaemia of approximately

108.1 trypanosomes/mL blood. The trypanosome strains used were Trypanosoma congolense

IL1180, a strain originating from Serengeti in Tanzania (Geigy and Kauffman, 1973) and T. b.

brucei ANTAR 1, a strain derived from the stock EATRO 1125 (Le Ray et al., 1977). Control

flies were given a bloodmeal on uninfected mice. Whole abdomens were removed from the

infected flies on days 1, 3 or 5 following the initial bloodmeal and pooled by two for total

RNA extraction.

5.2.5. RNA extraction and quantification

Immediately after removal, the abdominal tissue was manually homogenized with a Teflon

pestle in 1 mL Tripure® reagent (Roche Diagnostics GmbH, Mannheim, Germany) and the

total RNA was extracted according to the manufacturer’s protocol. To remove any

contaminating genomic DNA, the total RNA extracts were treated with DNase I (DNA-freeTM;

Ambion Inc., Austin, UK). Then, the RNA content was quantified using a Nanodrop®


66

spectrophotometer (NanoDrop Technologies Inc., Wilmington, DE, USA). The total RNA

extracts were stored at -20° until further use.

5.2.6. First- strand cDNA synthesis

Total RNA (400 ng) was mixed with 100 pmol Oligo(dT)15 Primer (Promega, Madison, WI,

USA) and RNAse free water to a total volume of 13.5 µL, incubated for 5 min at 65°C and

immediately cooled on ice for 1 min. Then, 4 µL Transcriptor reverse transcriptase (RT)

reaction buffer (Roche, Diagonstics GmbH), 2 µL dNTP-mix (10 mM each) and 0.5 µL

Transcriptor RT (10 units) (Roche, Diagonstics GmbH) were added. The reaction mixture was

incubated for 30 min at 55°C, followed by 5 min at 85°C and finally chilled on ice.

5.2.7. Quantitative Real - time PCR

Quantitative Real-time polymerase chain reaction (PCR) was performed in a 25µL PCR-

reaction mixture that contained 1 µL of the primary cDNA, 12.5 µL of iQ SYBR Green

Supermix (Bio-Rad Laboratories, Hercules, CA, USA) and an optimalized concentration of a

primer pair for one of the immunopeptide genes attacin (300 nM), defensin (700 nM) or

cecropin (300 nM). The following primer pairs were used: attacinFW: 5’-

TTTTCACAGTCGCACCCATT-3’ and attacinREV: 5’-AAACGCCTCCTGTCAAATCC-3’,

defensinFW: 5-TAGTTTTGGCTTTTCTTACAC-3’; defensinREV: 5’-

CGACTACAGTATCCGCTCTTT-3’ and cecropinFW: 5’-ATACTCGCTCTTTCAGTCAG-

3’ and cecropinREV: 5’-CTCTAACAGTAGCGGCAACA-3’. For the amplification of the

“housekeeping” genes actin (700 nM) and tubulin (300 nM) the following primer pairs were

used: actinFW: 5’-CGCTTCTGGTCGTACTACT-3’ and actinREV: 5’-

CCGGACATCACAATGTTGG-3’, tubulinFW: 5’-GATGGTCAAGTGCGATCCT-3’ and

tubulinREV: 5’-TGAGAACTCGCCTTCTTCC-3’. Reactions were run in an iCycler iQ

detection system (Bio-Rad Laboratories) and analysed with its software Version 3.1. The PCR

conditions comprised initial 10-min polymerase activation at 95°C followed by 35 cycles,

each consisting of a denaturation step at 95°C for 15 s and an annealing/elongation step at

60°C for 60 s. Three ‘technical’ replicates were performed for each sample and threshold

cycles (Ct) were recorded and used to calculate gene expression levels.


All statistical analyses were carried out in STATA Version 9.2 (StataCorp, Inc., College

Station, TX, USA). A linear regression was used to analyse loss of weight in individual flies.

The weight difference was used as the response and fly groups (starved and non-starved

teneral and adult flies) as the discrete explanatory variable. The effect of starvation on


67

expression levels of attacin, defensin and cecropin in teneral and adult flies was analysed

separately using a robust linear model. The response variables were the logarithm of the

normalized number of cycles needed for the immunopeptides specific cDNA to reach the

threshold in real-time PCR. The normalization used was a modification of that proposed by

Vandesompele et al. (2002):

( )[ ] ( )[ ] ( )[ ]2

lnlnln

tubact

pep

Ct

tub

Ct

actCt

pep

PCRPCRPCRresponse

+−=

with:

• PCRpep/act/tub = PCR efficiency of immunopeptides, actin and tubulin respectively

• Ctpep = number of cycles required for immune peptides genes to reach the threshold in

each sample repetition and

• Ctact/tub = average number of cycles (from three repetitions) required for actin and

tubulin genes, respectively, to reach the threshold in each sample.

Clustering effects resulting from repeated measures of the same samples were taken into

account. Discrete explanatory variables consisted of the different experimental groups created

in each experiment. Differences were considered significant when P < 0.05.

5.3. Results

5.3.1. Fat content and nutritional stress

Non-starved 20-day-old flies (AD2) had an approximately 3-fold higher fat content compared

with freshly emerged flies (TF0) (6.9 ± 0.2 mg vs 2.5 ± 0.2 mg). After a period of starvation

of four or seven days in teneral and 20-day-old flies, respectively, a 3-fold decrease in fat

content was observed (P < 0.0001). Starving 20-day-old flies for seven days (AD7) reduced

their fat content to a level similar to that in non-starved teneral flies (TF0) (Figure 5.1).

Experimental group

TF0 TF4 AD2 AD7

Fat

con

ten

t (m

g/f

ly)

0

2

4

6

8

***

***

Figure 5.1: Fat content (± 95% confidence intervals; n=12 per group) of male teneral and 20-day-old G. m. morsitans after a period of starvation. The fat content was significantly reduced in teneral starved 4 days (TF4) and 20-day-old flies starved 7 days (AD7) (*** p<0.0001) compared with non-starved teneral (TF0) and 20-day-old (AD2) flies.


68

5.3.2. Immune gene expression level and nutritional stress

Uninduced baseline levels of immune gene expression in non-starved teneral and 20-day-old

flies were similar except for that of defensin, where expression was 10-fold higher in the older

flies. Starving a teneral tsetse fly for four days significantly reduced its baseline levels of

expression of attacin and cecropin (compared with non-starved teneral flies), whereas the

observed decrease in defensin expression was not statistically not significant (P = 0.07)

(Figure 5.2A). In 20-day-old flies, starvation did not significantly compromise baseline

expression levels of these three immune genes (Figure 5.2B).

Experimental groupTF0 TF4

Norm

aliz

ed e

xpre

ssio

n v

alues

0,1

1

10

Experimental group

AD2 AD7

0,1

1

10

***

**

A. B.

Figure 5.2: Normalized expression levels (± 95% confidence intervals; n=12 per group) of attacin

(dark grey bar), defensin (white bar) and cecropin (light grey bar) in male teneral and 20-day-old

G. m. morsitans after starvation. Expression levels (mRNA) of attacin, defensin and cecropin

were measured in non-starved (TF0) and starved (TF4) teneral (A) and non-starved (AD2) and

starved (AD7) 20-day-old flies (B) using quantitative real-time PCR. Immunopeptide gene

expression values were normalized against the expression levels of the housekeeping genes actin

and tubulin. The expression levels of attacin and cecropin were significantly reduced in starved

teneral starved for four days (TF4) compared with non-starved teneral flies (TF0) (*** p<0.0001;

** p<0.02).


69

5.3.3. Immune gene response to bacterial/trypanosomal challenge and nutritional stress

Injection of bacteria (E. coli) induced high levels of expression of the genes in all

experimental groups. No mortality of flies was observed after the injection. In the teneral flies

(Figure 5.3A), the bacteria-induced increases in immune gene expression were > 100-fold for

the attacin and cecropin genes and around 40-fold for the defensin gene greater than those in

control flies. No significant differences were observed between the non-starved and starved

flies challenged with bacteria. In the 20-day-old flies (Figure 5.3B), the bacteria-induced

increase in immune gene expression was around 400-fold for attacin and defensin and 1000-

fold for cecropin greater than in control flies. Again, no significant difference in this immune

response was observed between starved and non-starved flies.

Experimental group

TF0+PBS TF0+E.coli TF4+PBS TF4+E.coli

No

rmal

ized

ex

pre

ssio

n v

alu

es

0,1

1

10

100

1000

10000

Experimental group

AD2+PBS AD2+E.coli AD7+PBS AD7+E.coli

0,1

1

10

100

1000

10000A. B.

******

*********

*********

***

******

***

Figure 5.3: Normalized expression levels (± 95% confidence intervals; n=4 per group) of attacin

(dark grey bar), defensin (white bar) and cecropin (light grey bar) in non- starved (TF0) and starved

(TF4) teneral (A) and in non-starved (AD2) and starved (AD7) 20-day-old (B) male G. m.

morsitans after bacterial (E. coli) challenge. The immunopeptide gene expression values were

normalized against the expression levels of the housekeeping genes actin and tubulin. Expression

levels of the three AMPs highly upregulated in the E.coli stimulated flies (***p<0.0001).


70

For the trypanosomal challenge, flies were given a bloodmeal containing T.

congolense or T. b. brucei bloodstream forms. Immune gene expression levels were

monitored for the first five days following the parasite uptake. Five days after ingestion of the

bloodmeal containing T. congolense or T. b. brucei, no increased expression levels of attacin,

defensin and cecropin genes were observed in teneral tsetse flies compared with those

observed on day 1 (Figures 5.4A and 5.5A). By contrast, in both non-starved and starved

teneral flies challenged with T. congolense (Figure 5.4A), gene expression levels of these

immunopeptides (except for defensin) on day 5 were significantly lower than on day 1.

Experimental group

AD2Tcd1 AD2Tcd5 AD7Tcd1 AD7Tcd5

0,01

0,1

1

10

100

Experimental group

TF0Tcd1 TF0Tcd5 TF4Tcd1 TF4Tcd5

Norm

aliz

ed e

xp

ress

ion

val

ues

0,01

0,1

1A. B.

***

******

**

Figure 5.4: Normalized gene expression levels (± 95% confidence intervals; n=5 per group)

of attacin (dark grey bar), defensin (white bar) and cecropin (light grey bar) in non-starved

(TF0) and starved (TF4) teneral (A) and non-starved (AD2) and starved (AD7) 20-day-old

flies (B) G. m. morsitans on day 1 (d1) or day 5 (d5) after uptake of a bloodmeal containing T.

congolense (Tc) bloodstream forms. Immunopeptide gene expression values were normalized

against the expression levels of the housekeeping genes actin and tubulin. The expression

levels of attacin and cecropin were significantly reduced in teneral starved for four days (TF4)

compared with non-starved teneral flies (TF0) (*** p<0.0001; ** p<0.03).


71

Non-starved 20-day-old tsetse challenged with T. congolense or T. b. brucei showed

increased levels of expression attacin, defensin and cecropin five days after the infective

blood meal compared to day 1 (Figures 5.4B and 5.5B), although this increase was

demonstrated as statistically significant only for the T. b. brucei-infected group. No

significant changes in immune gene expression levels were observed in the starved 20-day-

old tsetse fly group after a bloodmeal containing T. congolense or T. b. brucei.

Experimental group

AD2Tbd1 AD2Tbd5 AD7Tbd1 AD7Tbd5

0,01

0,1

1

10

100

Experimental group

TF0Tbd1 TF0Tbd5 TF4Tbd1 TF4Tbd5

No

rmal

ized

ex

pre

ssio

n v

alues

0,01

0,1

1

***

A. B.

***

***

**

Figure 5.5: Normalized gene expression levels (± 95% confidence intervals; n=5 per group) of

attacin (dark grey bar), defensin (white bar) and cecropin (light grey bar) in non-starved (TF0) and

starved (TF4) teneral flies (A) and non-starved (AD2) and starved (AD7) 20-day-old flies (B) G. m.

morsitans on day 1 (d1) or day 5 (d5) after uptake of a bloodmeal containing T. brucei brucei (Tb)

bloodstream forms. Immunopeptide genes expression values were normalized against the

expression levels of the housekeeping genes actin and tubulin. The expression levels of the

immunopeptide genes were significantly upregulated in non-starved adult flies after a bloodmeal

with T. brucei bloodstream forms (*** p<0.0001; ** p<0.03; * p=0.06).


72

5.4. Discussion

Trypanosome transmission is compromised by a natural tsetse fly refractoriness to

trypanosome infection because only a small subset of flies allow the development of a mature

infection in the hypopharynx (T. congolense) or salivary glands (T. brucei). One of the

defence mechanisms implicated in this refractoriness is the tsetse fly’s immune response. An

important part of this immune response is based upon the expression of several AMPs that are

mainly produced systemically by the fat body (Hao et al., 2001; Boulanger et al., 2002). In

addition, it was shown that the nutritional status of the tsetse fly at the time of the infective

bloodmeal affects its vectorial ability for both T. congolense and T. brucei brucei. Indeed, an

extreme period of starvation (four days for freshly emerged flies, seven days for older flies)

lowers the developmental barrier for trypanosome infection, especially at the midgut level of

the fly (Kubi et al., 2006). Hence, in this study, the effect of starvation (= nutritional stress)

was evaluated on the uninduced baseline levels of gene expression of the AMPs attacin,

defensin and cecropin in the tsetse fly and their induced levels of expression in response to

bacterial (E. coli) or trypanosomal challenge. The injection of bacteria causes a direct and

strong stimulation of the tsetse fat body immune response (Hao et al., 2001; Boulanger et al.,

2002) and is an appropriate assay to evaluate whether the responsiveness of the fly’s major

immune responsive organ is affected by nutritional deprivation. Ingestion of trypanosome

parasites in the tsetse midgut is also reported to indirectly induce a differential fat body

response, but the exact nature of this trypanosome-related stimulus is not yet clearly

understood (Lehane et al., 2008). In addition, local production of specific AMPs in the tsetse

midgut in response to trypanosome parasites cannot be excluded as the gut of a range of

insects was found to be a site of AMP synthesis (Boulanger et al., 2006). However, the AMP

gene expression analysis performed in this study did not allow us to make this distinction

between local and systemic immune response because it was based on RNA that was

extracted from the whole fly abdomen containing both the fat body and the tsetse midgut.

Both freshly emerged flies and 20-day-old G. m. morsitans flies showed significant

baseline gene expression of all three AMPs with a 10-fold increased in defensin expression

observed in the 20-day-old flies. However, starving freshly emerged flies for four days to a

fat reserve level of < 1 mg/fly significantly reduced this level of immune gene expression.

This suggests that high nutritional stress in these young flies results in a significant reduction

in uninduced baseline immune gene expression (i.e. innate expression before acquisition of

trypanosomes), which in turn, may contribute to the increased susceptibility of these flies to


73

trypanosome infection. Indeed, a correlation between this baseline immune gene level and the

fly’s susceptibility to trypanosome infection was recently suggested by Nayduch and Aksoy

(2007), who showed that the uninduced baseline level of systemic attacin was significantly

higher in freshly emerged flies of trypanosome-refractory tsetse species than in susceptible

species. In 20-day-old flies, the effects of nutritional stress by starvation were less pronounced

(i.e. a 3-fold reduction in fat level, which is comparable with that in non-starved teneral flies),

which allowed the flies to maintain a baseline level of immunopeptide expression similar to

those in non-starved flies.

A bacterial challenge by live E. coli injection resulted in high increases in expression of

attacin, defensin and cecropin in all experimental fly groups, thus confirming the strong

immunogenic nature of an E. coli injection to tsetse flies (Hao et al., 2001; Boulanger et al.,

2002; Lehane et al., 2004). Flies aged 20 days showed a much higher degree of

responsiveness to the bacterial challenge than freshly emerged flies, with a 400-fold induction

for attacin and defensin and a 1000-fold induction for cecropin in comparison with uninduced

baseline levels. These data clearly show that the fat body, as the major immune response

organ, is operational in freshly emerged, unfed flies, but has not yet attained its full capacity

to respond to pathogens. Moreover, as no differences were found between the starved and

non-starved groups, it is clear that this immune responsiveness to the bacterial challenge is

not affected by the nutritional status of the flies. This indicates that, although the maintenance

of a powerful immune defence system represents a high cost in energy for the tsetse fly

(Schmid-Hempel, 2005), keeping a properly functioning and alert immune defence system is

of vital importance to the fly, even when its energy reserve is seriously depleted. The uptake

of approximately 5 x 106 bloodstream trypanosomes (T. congolense or T. b. brucei) by blood-

feeding did not affect expression levels of attacin, defensin and cecropin in the teneral fly

groups at five days after the infective bloodmeal, which confirms previous observations by

Hao et al. (2001) that the immune system response to ingested bloodstream trypanosomes in

young tsetse flies is low. In the adult fly group, gene expression of attacin, defensin and

cecropin in response to trypanosome infection (especially for T. b. brucei) increased

significantly only in non-starved flies; this may represent a contributing factor to the high

refractoriness for trypanosome infection observed in these flies (Kubi et al., 2006).

In conclusion, this study reports that high nutritional stress decreases the baseline

immunopeptide gene expression in newly hatched unfed tsetse flies and decreases the immune

responsiveness of older flies to trypanosome challenge. This decreased immune gene

expression as a result of starvation may contribute to the increased susceptibility of


74

nutritionally stressed tsetse flies developing a trypanosome infection. It is clear that the

present study is based only on a limited experimental read-out of three well-characterized

immune-responsive genes in tsetse (attacin, defensin and cecropin), regulated by the IMD

pathway, which are assumed to represente a barometer for the functioning of the fat body-

regulated immune response. It would be worthwhile expanding this starvation study to the

expression of other fat body-regulated genes that are differentially expressed in trypanosome-

challenged tsetse flies.


75

5.5. References

Abubakar, L.U., Bulimo,W.D., Mulaa,F.J., Osir,E.O., 2006. Molecular characterization of a tsetse fly midgut proteolytic lectin that mediates differentiation of African trypanosomes. Insect Biochemistry and Molecular Biology 36: 344-352.

Aksoy, S., Gibson, W.C., Lehane, M.J., 2003. Interactions between tsetse trypanosomes with implications for the control of trypanosomiasis. Advances in Parasitology 53:1-83

Attardo, G.M., Strickler-Dinglasan, P., Perkin, S.A.H., Caler, E., Bonaldo, M.F., Soares, M.B., El-Sayeed, N., Aksoy, S., 2006. Analysis of fat body transcriptome from the adult tsetse fly, Glossina morsitans morsitans. Insect Molecular Biology 15: 411-424

Boulanger, N., Brun, R., Ehret-Sabatier, L., Kunz, C., Bulet, P., 2002. Immunopeptides in the defense reactions of Glossina morsitans to bacterial and Trypanosoma brucei brucei infections. Insect Biochemistry and Molecular Biology 32: 369-375

Boulanger, N., Bulet, P., Lowenberger, C., 2006. Antimicrobial peptides in the interactions between insects and flagellate parasites. Trends in Parasitology 22 : 262-268

Geigy, R., Kauffman, M., 1973. Sleeping sickness survey in the Serengeti area (Tanzania) 1971. I. Examination of large mammals for trypanosomes. Acta Tropica 30: 12-23

Gingrich, J.B., Ward, R.A., Macken, L.M., Esser, K.M., 1982. African sleeping sickness: new evidence that mature tsetse flies (Glossina morsitans) can become potent vectors. Transactions of the Royal Society of Tropical Medicine and Hygiene 76: 479-481


Sciences of the USA 98:12648-12653

Hu, C., Aksoy, S., 2006. Innate immune responses regulate trypanosome parasite infection of the tsetse fly Glossina morsitans morsitans. Molecular Microbiology 60: 1194-1204

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Van den Bossche, P., 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392

Langley, P.A., Hargrove, J.W., Wall, R.L., 1990. Maturation of the tsetse fly Glossina pallidipes (Diptera: Glossinidae) in relation to trap-orientated behaviour. Physiological

Entomology 15: 179-186

Leak, S.G.A., 1999. Tsetse biology and ecology: Their role in the epidemiology and control of trypanosomosis. CABI Publishing/ International Livestock Research Institute (ILRI), Wallingford, 568pp.



76

Lehane,M.J., Gibson,W., Lehane S.M., 2008. Differential expression of fat body genes in Glossina morsitans morsitans following infection with Trypanosoma brucei brucei. International Journal for Parasitology 38: 93-101

Le Ray, D., Barry, J.D. Easton, C., Vickerman, K., 1977. First tsetse fly transmission of the ‘Antat’ serodeme of Trypanosoma brucei. Annales de la Société Belge de Médecine

Tropicale 57 : 369-381

Macleod, E.T., Maudlin, I., Darby, A.C., Welburn, S.C., 2007. Antioxidants promote establishment of trypanosome infections in tsetse. Parasitology 134: 827-831.

Munks, R.J., Sant’Anna, M.R., Grail, W., Gibson, W., Igglesden, T., Yoshiyama, M., Lehane, S.M., Lehane, M.J., 2005. Antioxidant gene expression in the blood-feeding fly Glossina morsitans morsitans. Insect Molecular Biology 14: 483-491

Nayduch, D., Aksoy, S., 2007. Refractoriness in Tsetse Flies (Diptera: Glossinidae) May be a Matter of Timing. Journal of Medical Entomology 44: 660-665

Schmid-Hempel, P., 2005. Evolutionary ecology of insect immune defenses. Annual Review

of Entomology 50: 529-551

Van Den Abbeele, J., Claes, Y., Van Bockstaele, D., Le Ray, D., Coosemans, M., 1999. Trypanosoma brucei spp. Development in the tsetse fly: characterization of the post-mesocyclic stages in the foregut and proboscis. Parasitology 118: 469-478

Vandesompele, J., De Preter, K., Pattyn, F., Poppe, B., Van Roy, N., De Paepe, A., Speleman, F., 2002. Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biology 3: Research 0034.1-0034.11

Chapter 6

Investigation of the effect of seasonal climatic change on

nutritional and immune status and trypanosome infection in

natural field-caught tsetse flies

Chapter 6: Seasonal climatic change and tsetse fly nutritional & immune status

78

6.1. Introduction

An important factor in the complex epidemiology of tsetse-transmitted trypanosomiasis is the

proportion of infected flies transmitting the disease. This proportion is affected by several

tsetse and trypanosome-related factors. The majority of tsetse flies are refractory to infections

with trypanosomes. Furthermore, the tsetse fly susceptibility to infection varies according to

the trypanosome species and is associated with the complexity of the trypanosome

developmental cycle in the fly (Leak, 1999). Trypanosoma vivax, having a simple cycle in

the tsetse’s mouthparts, is easily transmitted whereas the complex cycle of T. brucei s.l.

results in low infection rates. Moreover, some tsetse species seem to be less susceptible to

trypanosome infections than others. Within a species, various extrinsic and intrinsic factors

have been described that affect the fly susceptibility to infection. It is generally accepted that

the majority of tsetse flies acquires a T. congolense or T. brucei spp. infection at the teneral

state during the first blood meal. Adult flies are considered to contribute less in trypanosome

transmission. However, Kubi et al. (2006) have shown that starvation significantly increases

the susceptibility of teneral and adult tsetse flies to trypanosomal infections. Similarly, Akoda

et al. (2009a) recently demonstrated that nutritional stress of reproducing female tsetse flies

resulted in a significant increase of their offspring’s vectorial capacity.

Various compounds associated with the tsetse immune system have been proven to

have an effect on the development and maturation of trypanosome infections in tsetse flies

(Hao et al., 2001; Boulanger et al., 2002; Hu and Aksoy, 2006). Attempts to elucidate the

mechanism involved in the changes in susceptibility to infection, showed a correlation

between susceptibility and the down-regulation of the fly’s immune response as a result of the

depletion of the fat body due to starvation (Akoda et al., 2009b). Under field conditions, tsetse

flies are subjected to considerable changes in their fat content (Glasgow and Bursell, 1960;

Van den Bossche and Hargrove, 1999). Especially during the hot dry season, fat body

reserves are generally low and body size decreases with smaller flies emerging as a result of

lower nutritional reserves in the mother. Whether such reductions in the fat body reserves also

results in a reduced expression of immune genes and increased susceptibility to trypanosomal

infections in flies captured in the field needs to be investigated further. For this purpose, a

study was undertaken to compare the expression of three immune peptides in a tsetse

population during the rainy and the hot dry season.


79


6.2.1. Study site

The field work was conducted at Rekomitjie Research Station in the Zambezi Valley,

Zimbabwe (16°18’S; 29°23’E, altitude 510m) (Figure 6.1) in April and October

corresponding respectively to the end of the rainy season (December to April) and the hot dry

season (September to November) (Gibson and Torr, 1999). In the Zambezi Valley, October

and November are the hottest months of the year with average maximum temperatures

ranging between 35 and 40°C and relative humidities of 40-60% (Gibson and Torr, 1999;

Torr and Hargrove, 1999; Muzari and Hargrove, 2005). Tsetse flies were collected in the

riverine woodland surrounding the Research Station. This area supports large populations of

game animals and a few cattle kept for research purposes. Both Glossina pallidipes Austen

and G. m. morsitans Westwood occur in the study area, although G. pallidipes is the more

abundant (Hargrove, 1981). Daily screen temperature and rainfall during the study period

were recorded at Rekomitjie Research Station.

Figure 6.1: A map of Zimbabwe showing the Zambezi Valley and the sampling area


80

6.2.2. Trapping of tsetse and sample collection

Tsetse flies were captured using fly rounds for G. m. morsitans and acetone baited-epsilon

trap for G. pallidipes. Flies were collected every morning between 07:00 and 09:00h and

dissected the same day to determine their infection status. For each of the two collection

periods (rainy and hot dry season), the abdomens of 50 and 20 males G. m. morsitans and G.

pallidipes, respectively, were preserved in RNAlater reagent (Ambion) to conserve the total

RNA for immune peptides expression analyses. Moreover, 100 male G. m. morsitans and 50

male G. pallidipes were killed immediately after capture and dried to determine their body fat

content.

6.2.3. Fly dissection

The non-teneral flies were dissected in a drop of sterile 0.9% saline solution. Mouthparts,

salivary glands and midgut of each fly were dissected using Lloyd and Johnson’s (1924)

method and successively examined under a light microscope (400x) to determine the presence

of trypanosomes. Positive mouthparts and/or midgut were spotted onto Whatman® FTA

cards, air-dried for one hour and stored at ambient temperature for future PCR analyses to

identify trypanosomes species.

6.2.4. Trypanosome species identification

A nested PCR-based method for trypanosome species identification (Adams et al., 2006) was

used to identify the trypanosome species in the positive field samples. Two mm diameter disc

of Whatman® FTA cards containing trypanosome-infected organs were prepared for PCR

analysis following the manufacturer’s instructions. PCR primers, amplification conditions,

and size analysis of the amplification products were performed as described in the

publication. As a modification, primary PCR products were enzyme-treated with ExoSAP-IT

to remove excess of dNTPs and the primary primers. One µL of a 1/25 dilution of this

primary product was used as template in the nested PCR-reaction. After the agarose gel

electrophoresis, a selection of amplified fragments were cloned into the plasmid vector PCR

2.1 (Invitrogen) and subsequently sequenced. BLAST analysis of the obtained sequences

allowed us to verify the link between the amplification fragment size and the trypanosome

species identification as was suggested by Adams et al. (2006).

6.2.5. Fat body content and immune peptide expression analysis

For a detailed description: See Chapter 5.


81


Statistical analyses to compare the fat body content and the expression levels of attacin,

defensin and cecropin in tsetse flies collected during the end of rainy and the hot dry seasons

were performed as described in chapter 5 section 5.2.8.

6.3. Results

6.3.1. Fat body content and immune peptide expression levels

The fat content in male G. m. morsitans collected during the hot dry season was significantly

lower compared to the fat content of flies captured during the end of the rainy season

(p<0.0001) (Figure 6.2). Moreover, the normalized expression levels of attacin, defensin and

cecropin were significantly lower in G. m. morsitans captured during the hot dry season

compared to expression levels at the end of the rainy season (Figures 6.3A). In G. pallidipes,

on the other hand, no significant difference in fat body content was observed in flies collected

during the two seasons. With the exception of defensin, the normalized expression of the

other two immune peptides was reduced significantly in G. pallidipes males collected during

the hot dry season (Figures 6.2 and 6.3B).

Figure 6.2: Fat body content (± 95% confidence intervals) of male non-teneral G. m.

morsitans (n= 100) and G. pallidipes (n=50) collected during a rainy season and hot dry

season in the field. The fat content was significantly reduced in G. m. morsitans flies collected

during the hot dry season (***p<0.0001) compared to the same species collected during the

rainy season. In G. pallidipes, fat body content did not differ between the two seasons of

collection.

0

0,5

1

1,5

2

2,5

3

3,5

G.m. morsitans G. pallidipes

Fat body c

onte

nt (m

g/fly

)

Rainy season

Hot dry season

***


82

Figure 6.3: Normalized expression levels (± 95% confidence intervals; n=30 per group) of

attacin, defensin and cecropin in male non-teneral G. m. morsitans (A) and G. pallidipes (B)

collected during the rainy season and the hot dry season in the Zambezi Valley of

Zimbabwe. Expression levels (mRNA) of attacin, defensin and cecropin were measured

using quantitative real-time PCR. The immune peptide gene expression values were

normalized against the expression levels of the housekeeping genes actin and tubulin.

Except for defensin in G. pallidipes, the expression levels of the immune peptides were

significantly reduced in flies collected during the hot season (***p < 0.0001; **p < 0.001)

A

B

Rainy season Hot dry season

No

rma

lize

d e

xp

ressio

n le

ve

ls

0.00

0.03

0.05

1.00

2.00 Attacin

Defensin

Cecropin

Rainy season Hot dry season

No

rma

lize

d e

xpre

ssio

n le

ve

l

0.00

0.05

0.10

0.15

5.00

10.00

15.00 Attacin

Defensin

Cecropin

***

***

** **

**


83

6.3.2. Tsetse infection rates and trypanosome species

A total of 449 (267 G. m. morsitans and 182 G. pallidipes) and 420 (204 G. m. morsitans and

216 G. pallidipes) non-teneral flies were dissected during the rainy season and the hot dry

season, respectively. A total of 6.3% (29/449) of the flies captured during the rainy season

and 9.3% (39/420) of the flies captured during the hot dry season, had infections in midgut

and/or proboscis or salivary glands when examined microscopically (Table 6.1).

Table 6.1: Number and proportion (%) of non-teneral (male and females) tsetse G. m.

morsitans and G. pallidipes infected with trypanosomes in the rainy season and the hot dry

season.

Number (%) of infected flies

Fly species Seasons

Total

dissected Midgut Proboscis Salivary glands

Rainy season 267 17 (6.3) 10 (3.7) 0 (0.0)

G.m.morsitans Hot dry season 204 16 (7.8) 17 (8.3) 0 (0.0)

Rainy season 182 6 (3.2) 10 (5.4) 1 (0.5)

G. pallidipes Hot dry season 216 8 (3.7) 14 (6.4) 1 (0.4)

Nested and standard PCR combined with sequence analyses of microscopically positive

midguts and proboscis revealed the presence of a diversity of different trypanosome species in

the flies such as T. vivax, T. congolense s.l, T. brucei s.l., T. simiae and T. godfreyi. Of the

positive midguts collected in hot dry season 54.1% (13/24) and 20.8% (5/24) had an infection

with T. congolense or T. brucei s.l. respectively. Of the positive midguts collected in rainy

season 52.2% (12/23) and 17.4% (4/23) had an infection with T. congolense or T. brucei s.l.

respectively.

6.4. Discussion

The present study aimed to prospect the effect of seasonal climatic change on the nutritional

status and the expression level of a limited selection of immune peptide genes in field-caught

tsetse flies.

The results showed that the nutritional status (measured by the fat content) of G.

pallidipes and G .m. morsitans was similar during the rainy season although G. pallidipes

flies are larger and have proportionally higher food reserve and fat content than G. m.

morsitans (Bursell, 1960; Phelps, 1973). However, in G. pallidipes, the nutritional status did

not vary with seasonal climatic change while the immune gene


84

expression levels were significantly affected. These observations are difficult to interpret

because of the bias associated to the used capture method. Indeed, the traps which were used

for G. pallidipe will only sample the hungriest elements of the tsetse population. By

consequence, the measured nutritional status of the captured flies underestimates the actual

nutritional status of the population (Hargrove, 1999). Hence, further research is needed to

investigate the role of seasonal climatic change in G. pallidipes nutritional and immune status

by using other sampling techniques such as artificial refuges which collect flies with a wider

range of nutritional status compared to other stationary baits (Phelps and Vale, 1978).

In G. m. morsitans, on the other hand, fad body content decreases significantly during

the hot dry season. This reduced fat content is in accordance with previous field observations

(Van den Bossche and Hargrove, 1999). This lower nutritional status might be due to a

reduced availability of hosts during the hot dry season and an increased metabolism as a result

of the high ambient temperature prevailing in the Zambezi Valley during the hot dry season.

Importantly, the expression level of each of the analyzed immune peptides was significantly

lower during the hot dry season. It thus seems that in accordance with laboratory observations

(Akoda et al., 2009b), a more stressful situation such as the environmental conditions during

the hot dry season affect the nutritional status of G. m. morsitans resulting in a down-

regulation of the expression of some immune genes produced in the fat body (Hao et al.,

2001; Boulanger et al., 2002).

Notwithstanding the observed changes in immune peptide expression, the

repercussions of these changes for the vectorial capacity of tsetse flies are difficult to assess.

This is especially so since the number of flies captured and dissected did not allow the

establishment of an age-infection prevalence relationship that would highlight changes in the

susceptibility of especially adult flies. From these very limited data there is no indication that

seasonal change affects the fly infection rates. However, several previous field studies have

shown a relationship between seasonal climatic change and trypanosome infection rates in

tsetse flies. For example, field data collected throughout a period of 36 months in eastern

Zambia indicated that the monthly prevalence of trypanosome infection in tsetse was

associated to the average fat levels of the month with the highest prevalence of infection

occurring during the hottest months of the year when fat body level was lowest (Van den

Bossche et al. unpublished work). On a continental scale, a review of surveys determining the

infection rates of savannah species of tsetse flies conducted over a period of 45 years also

showed a positive correlation between infection rate of tsetse flies and the mean annual

temperature (Ford and Leggate, 1961).


85

In conclusion, this study demonstrates that the seasonal changes in the nutritional

status and immune peptide expression level do occur in field tsetse G m. morsitans

populations. However, the possible repercussions of such variation in trypanosome

transmission by this tsetse species require further investigations with larger samples.


86

6.5. Reference

Adams, E.R., Malele I.I., Msangi, A.R. and Gibson, W.C. 2006. Trypanosome identification in wild tsetse populations in Tanzania using generic primers to amplify the ribosomal RNA ITS-1 region. Acta Tropica 100: 103-109

Akoda, K., Van Den Abbeele, J., Marcotty, T., De Deken, R., Sidibe, I., Van den Bossche, P., 2009a. Nutritional stress of adult female tsetse flies (Diptera: Glossinidae) affects the susceptibility of their offspring to trypanosomal infections. Acta Tropica 111: 263–267

Akoda, K., Van den Bossche, P., Marcotty, T., Kubi, C., Coosemans, M., De Deken, R., Van Den Abbeele, J., 2009b. Nutritional stress affects the tsetse fly’s immune gene expression. Medical and Veterinary Entomology 23: 195-201


Bursell, E., 1960. The effect of temperature on the consumption of fat during pupal development in Glossina. Bulletin of Entomological Research 51: 583-598


Society of Tropical Medicine and Hygiene 55, 383-397.

Gibson, G., Torr, J., 1999. Visual and olfactory responses of haematophagous Diptera to host stimuli. Medical and Veterinary Entomology 13: 2-23

Glasgow, J. P. and Bursell, E., 1960. Seasonal variations in the fat content and size of Glossina swynnertoni Austen. Bulletin of Entomological Research 51: 705-713.

Hao, Z.R., Kasumba, I., Lehane, M.J., Gibson, W.C., Kwon, J., Aksoy, S., 2001. Tsetse immune responses and trypanosome transmission: Implications for the development of tsetse-based strategies to reduce trypanosomiasis. Proceedings of the National Academy

of Science of the USA 98: 12648-12653.

Hargrove, J.W., 1999. Lifetime changes in the nutritional characteristics of female tsetse Glossina pallidipes caught in odour-baited traps. Medical and Veterinary Entomology 13: 165-176

Hargrove, J.W., 1981. Discrepancies between estimates of tsetse fly populations using mark-recapture and removal trapping techniques. Journal of Applied Ecology 50: 351-373

Hu, C., Aksoy, S., 2006. Innate immune responses regulate trypanosome parasite infection of the tsetse fly Glossina morsitans morsitans. Molecular Microbiology 60:1194-1204.

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Van den Bossche P., 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392.


87

Leak, S.G.A., 1999. Tsetse biology and ecology: Their role in the epidemiology and control of trypanosomosis. CABI Publishing/ International Livestock Research Institute (ILRI), Wallingford, 568pp.

Llyod, L., Johnson, W.B., 1924. The trypanosome infections of tsetse flies in northern Nigeria and a new method of estimation. Bulletin of Entomological Resesearch 14: 265-288.

Muzari, M.O., Hargrove, J.W., 2005. Artificial larviposition sites for field collections of puperia of tsetse flies Glossina pallidipes and G. m. morsitans. (Diptera: Glossinidae). Bulletin of Entomological Research 95: 221-229

Phelps, R.J., 1973. The effect of temperature on fat consumption during the puparial stages of Glossina morsitans morsitans Westw. (Diptera: Glossinidae) under laboratories conditions and its implications in the field. Bulletin of Entomological Research 62: 423-438

Phelps, R.J., Vale, G.A., 1978. Studies on populations of Glossina morsitans morsitans and G. pallidipes (Diptera: Glossinidae) in Rhodesia. Journal of Applied Ecology 15: 743-760

Torr, S.J., Hargrove, J.W., 1999. Behavious of tsetse (Diptera: Glossinidae) during the hot season in Zimbabwe: the interaction of micro-climate and reproductive status. Bulletin

of Entomological Research 89: 365-379

Van den Bossche, P., Hargrove, J.W., 1999. Seasonal variation in nutritional levels of male tsetse flies Glossina morsitans morsitans Westwood (Diptera: Glossinidae) caught using fly-rounds and electric screens. Bulletin of Entomological Resesearch 89: 381-387.

Chapter 7

Investigations on the transmissibility of Trypanosoma congolense

by the tsetse fly Glossina morsitans morsitans during its

development in a mammalian host

Akoda K., Harouna S., Marcotty T., De Deken R. and Van den Bossche P. (2008).

Investigations on the transmissibility of Trypanosoma congolense by the tsetse fly Glossina

morsitans morsitans during its development in a mammalian host. Acta Tropica 107, 17-19

Chapter 7: T. congolense developmental stage in the host & transmissibility

89

7.1. Introduction

African trypanosomes causing sleeping sickness in humans and “Nagana” in livestock are

cyclically transmitted by tsetse fly, Glossina spp. Hence, the proportion of infected tsetse flies

in a population is of considerable importance in the epidemiology of human and animal

trypanosomiasis. Fly-related factors that affect the proportion of infected tsetse flies are well

documented (Kubi et al., 2006; Aksoy et al., 2003). Trypanosomes are also known to undergo

substantial metabolic changes to allow adaptation to the changing environment in the

mammalian host or insect vector (reviewed by Matthews, 2005). In polymorphic trypanosome

species such as T. brucei s.l., considerable morphological and metabolic changes prepare the

trypanosome for its survival and development in the tsetse midgut environment that is low in

glucose and oxygen (reviewed by Seed and Wenck, 2003). Therefore, the proportion of tsetse

flies in which a T. brucei infection develops is a function of the numbers of short stumpy

forms of the parasite in the blood meal (Wijers and Willett, 1960). It thus seems that for T.

brucei s.l., parasite-associated processes prepare the parasite for its development in the tsetse

fly and determines the ease with which the trypanosome is transmitted by tsetse flies.

Although T. congolense is generally considered to be monomorph, similar pleomorphism has

been described in some strains (Godfrey, 1960; Nantulya et al., 1978a) and has been

associated with the parasite’s transmissibility (Nantulya et al., 1978b). Nevertheless, with the

exception of the work described by Nantulya et al. (1978b), little is known of the

transmissibility of T. congolense during its development in the host. Here we investigate the

transmissibility of a monomorphic T. congolense strain during its development in a vertebrate

host.


7.2.1. Tsetse flies

Teneral male tsetse flies Glossina morsitans morsitans Westwood (less than 32 hours old)

from the colony maintained at the Institute of Tropical Medicine (Antwerp, Belgium) were

used in the experiments. The origin of this tsetse colony and the conditions of maintenance

are described by Elsen et al. (1993). This line of tsetse has a high vectorial capacity (Van Den

Abbeele, 2001).


90

7.2.2. Trypanosome

Trypanosoma congolense IL 1180, a strain originating from Serengeti in Tanzania (Geigy and

Kauffmann, 1973) was used for the infections. One cryostabilate of this strain was reactivated

in 3 mice (strain OF1) that served for experimental mice passage.

7.2.3. Experimental design

Each mouse of a pool of 25 mice (strain OF1) was injected with 0.2 ml of blood containing

106.9 trypanosomes per millilitre. From the third day after the infection onwards, the

parasitaemia of each mouse was measured daily using the scale of Herbert and Lumsden

(1976). On days 4, 5, 6, 7 or 10 post-infection, 4 mice were selected randomly from the pool.

The selected mice were anesthetized with a mixture of Ketalar® and Rompun® and one cage

containing 40 teneral male tsetse flies was placed on each anesthetized mouse to allow the

flies to take an infective blood meal. After the infective bloodmeal, flies which did not feed

were removed from the experiment. Engorged flies were retained and fed on clean rabbits 3

times a week until 48 hours before dissection. To avoid re-infection of the flies during the in

vivo maintenance, rabbits were replaced at weekly intervals. All surviving flies were dissected

21 days after the infective blood meal using the method described by Lloyd and Johnson

(1924). The midgut and mouthparts were examined for the presence of trypanosomes.

The proportion of immature (midgut) infections was calculated as the proportion of

dissected flies that had a trypanosome infection in the midgut. The maturation rate was

calculated as the proportion of midgut infection that developed into mature infection in the

mouthparts. The overall infection rates were calculated as the Intrinsic Vectorial Capacity

(IVC) described by Le Ray (1989). Statistical analyses of data were carried out in STATA

Version 9.2 (StataCorp, Inc., College Station, TX, USA) using a logistic regression to

compare proportions of immature and IVC in different experimental fly groups. Discrete

explanatory variables were the days following mice infection, the parasitaemia of the mouse

on which the infective blood meal was taken and the interaction between the two. Simplified

models (from which non-significant explanatory variables (parasitaemia) were dropped) were

selected when the likelihood ratio test had a P value > 0.05. Cluster effects resulting from

flies infected on the same mouse and maintained in the same cage were taken into account in

simplified robust models.


91

7.3. Results

Of a total of 800 male teneral flies, 70.5% (564/800) took the infective bloodmeal and were

retained for the experiment. The mortality rate was 0.9%. A total of 559 tsetse flies were

dissected to determine their infection status. The proportion of infected flies was significantly

higher (P < 0.05) in teneral tsetse flies infected on day 5 or 10 post-infection compared to the

proportion of infected flies that received an infective bloodmeal on the first day that parasites

were observed in the blood of the mice (day 4 post-infection) (Tables 7.1 and 7.2).

Proportion of infected flies

Mouse

Day

post-

infection

Parasitaemia

(antilog)

Number

dissected Midgut (p) Maturationa (m)

IVCb

1 4 6.9 35 0.20 (7/35) 1.00 (7/7) 0.20

2 4 6.9 29 0.34 (10/29) 1.00 (10/10) 0.34

3 4 < 5.4 33 0.27 (9/33) 1.00 (9/9) 0.27

4 4 6.9 26 0.27 (7/26) 0.86 (6/7) 0.23

5 5 7.8 35 0.51 (18/35) 0.89 (16/18) 0.46

6 5 6.9 25 0.48 (12/25) 0.92 (11/12) 0.44

7 5 8.1 30 0.40 (12/30) 1.00 (12/12) 0.40

8 5 7.2 25 0.44 (11/25) 1.00 (11/11) 0.44

9 6 8.4 30 0.10 (3/30) 1.00 (3/3) 0.10

10 6 8.4 30 0.23 (7/30) 1.00 (7/7) 0.23

11 6 8.1 19 0.47 (9/19) 1.00 (9/9) 0.47

12 6 8.1 29 0.41 (12/29) 0.92 (11/12) 0.38

13 7 8.1 26 0.38 (10/26) 1.00 (10/10) 0.38

14 7 8.4 31 0.13 (4/31) 1.00 (4/4) 0.13

15 7 8.4 25 0.44 (11/25) 1.00 (11/11) 0.44

16 7 8.1 28 0.36 (10/28) 0.80 (8/10) 0.29

17 10 6.9 28 0.57 (16/28) 1.00 (16/16) 0.57

18 10 6.9 26 0.46 (12/26) 1.00 (12/12) 0.46

19 10 8.1 27 0.48 (13/27) 1.00 (13/13) 0.48

20 10 7.2 22 0.59 (13/22) 1.00 (13/13) 0.59


mouthparts


Table 7.1: Immature and mature infection rates of male G. m. morsitans given a single

infective bloodmeal on various days of the development of T. congolense IL1180 in mice.


92

P values

Day post-infection Immature infection IVC

5 < 0.001 < 0.001

6 0.811 0.813

7 0.471 0.559

10 < 0.001 < 0.0001

At the peak of the parasitaemia (around day 7 post-infection), immature and mature

infection rates were not significantly different from the infection rates observed at the onset of

the parasitaemia (day 4 post-infection) (Tables 7.1 and 7.2 and Figure 7.1 for mature

infection).

Figure 7.1: Average proportion of mature infection of male G. m. morsitans infected on

different days post-infection on mice infected with T. congolense IL1180.

0

0,2

0,4

0,6

0,8

4 5 6 7 10

Day post infection

Matu

re in

fecti

on

pro

po

rtio

n

0

1

2

3

4

5

6

7

8

9

Para

sit

aem

ia (

An

tilo

g)

Mature infection Parasitaemia

Table 7.2: Significance (P value) of the difference between the proportion of male G. m. morsitans

with a mature or immature infection and infected on days 5, 6, 7 or 10 post-infection compared to

flies infected on day 4 post-infection (reference)


93

The effect of parasitaemia on the mature and immature infection rates was not

significant (p = 0.130 for immature and p = 0.281 for mature infections). The maturation rate

of the midgut infection was high and not affected by the day of infection (Table 6.1). The

design effect, reflecting the importance of intra-cluster correlation was small (DEFT range

between 0.55 and 1.25) indicating that individual mouse used to infect the flies had little

effect within experimental groups (day-post infection).

7.4. Discussion

Trypanosomes undergo a complex life cycle between the mammalian host and the insect

vector. The mechanisms of refractoriness or susceptibility of tsetse flies to a trypanosome

infection are not fully understood but parasite-related factors do play an important role in

determining the transmissibility of the trypanosome.

Contrary to earlier observations by Nantulya et al. (1978b), high transmissibility of T.

congolense does not seem to be associated with high parasitaemias. It is difficult to compare

both experiments since the experimental setup of Nantulya et al. (1978b) does not allow for a

thorough analysis of variability between fly batches and variability between days on the

rising, plateau or declining phase of the parasitaemia. Moreover, compared to Nantulya et al.

(1978a) we did not observe the morphological changes in the parasite during its development.

Our results show that the transmissibility of the T. congolense strain used was not

associated with differences in the parasitaemia of the mice at the moment of infection. Indeed,

the infection rate of tsetse infected on days 6 or 7 of the infection is significantly reduced even

when the parasitaemia is highest. This could be attributed to the effect of the host’s immune

system reducing the trypanosome’s viability and, hence their capacity to infect tsetse flies

(Morrison et al., 1985). On the other hand, the absence of a correlation between the infection

rate of tsetse and the parasitaemia at the moment of infection may not be surprising

considering the fact that a single trypanosome is sufficient to infect a tsetse fly (Maudlin and

Welburn, 1989). In polymorphic T. brucei s.l. also, transmissibility is not correlated with the

parasitaemia but with the proportion of non-dividing stumpy forms that constitute the

predominant population during the remission and relapse of the parasitaemia (Balber, 1972;

Van den Bossche et al., 2005). Such stumpy forms have the capacity to differentiate into

procyclic forms either in the tsetse midgut (Matthews, 1999) or in vitro (Breidbach et al.,

2002) whereas long slender forms are not fit to survive in the insect vector and die rapidly


94

after ingestion by the tsetse fly (Turner et al., 1988). Although such morphological changes

are not observed in the monomorphic T. congolense strain used in this experiment, metabolic

changes in the trypanosome could occur at each experimental stage and probably affect its

capacity to infect tsetse flies. Such metabolic changes could make the trypanosome less

susceptible to the elimination process in tsetse midgut immediately after the blood meal

ingestion (Van Den Abbeele et al., 1999). Although further research is required to determine

the processes involved in adapting bloodstream forms of T. congolense to the tsetse fly’s

midgut environment, the observed differences in the trypanosome’s transmissibility during its

development in the mammalian host stress the importance of standardising trypanosome

transmission experiments or experiments comparing the vectorial capacity of tsetse flies.

Furthermore, it would be interesting to confirm these finding using infected bovines.


95

7.5. References

Aksoy, S., Gibson, W.C., Lehane, M.J., 2003. Interactions between tsetse trypanosomes with implications for the control of trypanosomiasis. Advance in Parasitology 53: 1-83.

Balber, A.E., 1972. Trypanosoma brucei: Fluxes of the Morphological Variants in Intact and X-irradiated Mice. Experimental Parasitology 31: 307-319

Breidbach, T., Ngazoa, E., Steverding, D., 2002. Trypanosoma brucei: in vitro slender-to-stumpy differentiation of culture-adapted, monomorphic bloodstream form. Experimental Parasitology 101: 223-230

Elsen, P., Van Hees, J., Delil, E., 1993. L’historique et les conditions d’élevage des lignées de glossines (Diptera, Glossinidae) maintenues à l’Institut de Médecine Tropicale Prince Léopold d’Anvers. Journal of African. Zoology 107: 439-449.

Geigy, R., Kauffmann, M., 1973. Sleeping sickness survey in the Serengeti area (Tanzania) 1971: examination of large mammals for trypanosomes. Acta Tropica 30: 12-23

Godfrey, D.G., 1960. Types of Trypanosoma congolense. I. Morphological differences. Annal

of Tropical Medicine and Parasitology 54: 428-438.

Herbert, W.J., Lumsden, W.H.R., 1976. Trypanosoma brucei. A rapid matching method for estimating the host’s parasitemia. Experimental Parasitology 40: 427-431.

Kubi, C., Van Den Abbeele, J., De Deken, R., Marcotty, T., Dorny, P., Van Den Bossche, P., 2006. The effect of starvation on the susceptibility of teneral and non-teneral tsetse flies to trypanosome infection. Medical and Veterinary Entomology 20: 388-392



Llyod, L., Johnson, W.B., 1924. The trypanosome infections of tsetse flies in northern Nigeria and a new method of estimation. Bulletin of Entomological Research 14: 265-288.

Matthews, K.R., 1999. Development in the differentiation of Trypanosoma brucei. Parasitology Today 15: 76-80

Matthews, K.R., 2005. The developmental cell biology of Trypanosoma brucei. Journal of

Cell Sciences 118: 283-290

Maudlin, I., Welburn, S.C., 1989. A single trypanosome is sufficient to infect a tsetse fly. Annal of Tropical Medicine and Parasitology 83: 431-433.

Morrison W. I., Murray M., Akol G. W. O., 1985. Immune responses of cattle to African trypanosomes. In: I.R.Tizard (ed.), Immunology and Pathogenesis of Trypanosomiasis, CRC Press Inc., Florida, USA, pp. 103-31

Nantulya, V.M., Doyle, J.J., Jenni, L., 1978a. Studies on Trypanosoma (Nannomonas) congolense. I. On the morphological appearance of the parasite in the mouse. Acta

Tropica 35: 329-337


96

Nantulya, V.M., Doyle, J.J., Jenni, L., 1978b. Studies on Trypanosoma (Nannomonas) congolense. II. Observations on the cyclical transmission on three field isolates by Glossina morsitans morsitans. Acta Tropica 35: 329-337

Seed R.J., Wenck M.A., 2003. Role of the long slender to short stumpy transition in the life cycle of the African trypanosomes. Kinetoplastid Biology and Disease 2: 3

Turner, C.M.R., Barry, J.D., Vickerman, K., 1988. Loss of variable antigen during transformation of Trypanosoma brucei rhodesiense from bloodstream to procyclic forms in the tsetse fly. Parasitology Research 74: 507-511.

Van Den Abbeele, J., 2001. Trypanosoma brucei sp. Development in the tsetse fly Glossina morsitans: a parasitological and molecular approach, PhD Thesis. Antwerp University, Antwerp, 121pp.

Van Den Abbeele, J., Claes, Y., Van Bockstaele, D., Le Ray, D., 1999. Trypanosoma brucei spp. Development in the tsetse fly: characterization of the post-mesocyclic stages in the foregut and proboscis. Parasitology 118: 469-478.

Van den Bossche, P., Ky-Zerbo, A. X., Brandt, J., Marcotty, T., Geerts, S., De Deken, R., 2005. The transmissibility of Trypanosoma brucei during its development in cattle. Tropical Medicine and International Health 10: 833-839.

Wijers, D.J.B., Willet K.C., 1960. Factors that may influence the infection rate of Glossina palpalis with Trypanosoma gambiense. II. The number and the morphology of the trypanosomes present in the blood of the host at the infected feed. Annal of Tropical


Chapter 8

General discussion

Chapter 8: General discussion

98

8.1. Introduction

Understanding the interactions between the parasite and the vector is essential to develop

effective control strategies that reduce disease transmission. Over the years, various factors

that affect the establishment and subsequent maturation of a trypanosome infection in tsetse

flies have been identified (reviewed by Aksoy et al., 2003). However, our knowledge on the

biological mechanism of tsetse-trypanosome interaction is still limited. Nevertheless, ongoing

research in the innate immune responses of tsetse flies is shedding some light on the

molecular basis of refractoriness or susceptibility to trypanosome infection.

The main objectives of the present thesis were to improve our knowledge on i) the

effect of nutritional stress in tsetse flies on the transmission of human and animal pathogenic

trypanosome species, and ii) the molecular basis of the modulation of the tsetse fly

susceptibility to trypanosomal infections as a result of nutritional stress. Finally, the research

presented in this thesis aimed at determining the effects of seasonal climatic changes on the

nutritional status and immune peptide expression in field-caught tsetse flies in order to

determine whether changes in environmental conditions affect the trypanosome transmission

dynamics in the field. In the following sections and based on the findings of the research

described in the preceding chapters, we outline the relevance of these findings and present

them in the wider context of nutritional stress and environmental stressors.

8.2. Tsetse starvation and trypanosome development

The outcome of studies presented in this thesis, show that the nutritional stress at the moment

of the infective meal increases the tsetse flies’ susceptibility to develop not only animal

pathogenic trypanosomes but also the human pathogenic species T. b. rhodesiense. Moreover,

nutritional stress experienced by reproductive females increases the ability of their offspring

to transmit trypanosome infections. Furthermore, our research showed that the effects of

nutritional stress not only affects the development of a trypanosomal infection in the midgut

but also boosts the proportion of T. b. brucei midgut infections that matures into infectious

metacyclic forms. All these findings suggest that under specific environmental conditions that

affect the nutritional status of the flies, the tsetse population’s susceptibility to trypanosomal

infections may change drastically. The repercussions of these observations for our

understanding of the epidemiology of human and animal trypanosomiasis are considerable.

Especially in the case of human trypanosomiasis where the proportion of infected tsetse flies

is usually very low, any factor resulting in nutritional stress in a tsetse fly may increase the


99

fly’s susceptibility to trypanosome infections and induce the occurrence of disease epidemics

or facilitate its spread. Although our results show that nutritional stress certainly is a factor

affecting susceptibility through a decrease of the immune gene expression and hence reducing

the tsetse’s immune responsiveness, many other factors affecting the tsetse’s susceptibility

may be present.

8.3. Factors inducing nutritional stress in tsetse flies

Tsetse flies are haematophagous insects with both males and females feeding on mammalian

vertebrate hosts (mainly Bovidae and Suidae) at 2-3 days intervals (Randolph et al., 1992). In

adult tsetse, each blood meal is mostly devoted to the production of fat which provides the

energy for flight activity and reproduction (Rogers and Randolph, 1978). Depriving the flies

of food or increasing their metabolism will result directly in the reduction of fat level which

might decrease the immune responsiveness of the flies to the trypanosome parasite. Several

factors may contribute to nutritional stress in field populations of tsetse flies (Figure 8.1).

Figure 8.1: Diagram of parameters affecting fat depletion and immunosuppression in tsetse flies

Fat body depletion Immunosuppression

Nutritional stress Host availability

Climate (temperature)

• Geography

• Elevation

• Seasonal change

• Climatic change

• Habitat fragmentation

• Human encroachment


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8.3.1. Ambient temperature (climate)

The metabolism of tsetse flies is temperature dependent. Hence, an increase in the ambient

temperature will result in an increase in the metabolism of the fly and may reduce the fat body

content and, hence, result in a decrease of the immune responsiveness of a fly to the

trypanosome parasite. Ambient temperature thus seems to be an important determinant

affecting the susceptibility of a tsetse population to trypanosomal infections. Ambient

temperature is determined by a range of factors including geographic ones such as location

(latitude) and elevation (altitude). Moreover, it has important temporal connotations including

season and climate change.

8.3.1.1. Geographical location (latitude)

Africa is a vast continent with a wide variety of climate regimes. The continent is

predominantly tropical but there are subtropical regions and deserts. At the same altitude and

within the tropical zone, the climate becomes cooler when approaching the equator. It thus

seems that there is a North-South temperature gradient throughout the tsetse belt of tropical

Africa. According to the results presented in this thesis one would expect higher infection

rates to change according to this gradient. Interestingly, Ford and Leggate (1961) studying the

relationship between the trypanosome infection rate of tsetse populations and their geographic

location found a positive relationship between infection rates and distance from the equator.

This correlation indicates higher infection rates in hotter areas. A decreased

immunocompetence of the tsetse flies living in these hotter areas might be one of the

contributing factors to explain the higher trypanosome infection rates in tsetse.

8.3.1.2. The altitude

Irrespective of a geographic location, ambient temperature is also affected by the altitude with

valleys being much hotter than plateau areas. Again, according to the results presented in the

thesis, this would suggest higher susceptibility of flies present in hot areas such as valleys.

This hypothesis may explain the high prevalence of T. b. rhodesiense, a trypanosome species

that is notoriously difficult to transmit, in valley areas with high ambient temperatures.

Indeed, when comparing the maximum ambient temperatures in villages outside T. b.

rhodesiense foci but where tsetse flies are present (38.4 ± 0.60°C) with the maximum ambient

temperatures in villages inside T. b. rhodesiense foci (40.0 ± 0.49°C), T. b. rhodesiense foci

are significantly hotter (P<0.001) suggesting again a possible effect of temperature on T. b.

rhodesiense transmission dynamics (Figure 8.2) (Van den Bossche et al., unpublished work).


101

Figure 8.2: Map of the satellite measured average maximum temperature (A) and location of

tsetse belts, populated places and T. b. rhodesiense historical foci (B)

Method: Daily 1 km resolution NOAA-AVHRR satellite images for the period January 2001 to December 2002 were downloaded from the NOAA website (http://www.class.noaa.gow) and archived and processed using the Avia-GIS AVHRR processing software (http://www.avia-gis.com/site.html). Monthly cloud free maximum ground temperature images were computed using the Price-index. For each month a continental monthly maximum temperature map was produced by mosaicking the obtained temperature maps. Finally the annual maximum temperature for each pixel was determined using the macro modeler facility in Idrisi. Information on the limits of the historical T. b. rhodesiense foci in Botswana, Kenya, Malawi, Mozambique, Namibia, Tanzania, Uganda, Zambia and Zimbabwe was obtained from Sleeping Sickness Unit of the World Health Organization (WHO). To restrict the analysis to areas where people are present, the comparison of the maximum temperature was restricted to populated places georeferenced in the Digital Chart of the World database (http://www.maproom.psu.edu/dcw/) in areas where tsetse flies are present. To discriminate between tsetse presence and absence, continental predictions from the PAAT-IS were used (Van den Bossche et al. unpublished work).

A

B


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8.3.1.3. Seasonality

Ambient temperatures in a locality vary greatly between seasons. These substantial seasonal

climatic variations are known to have significant effects on the nutritional status of the tsetse

population represented by changes in the fat body content, the size of adult tsetse flies

(Glasgow and Bursell, 1960; Van den Bossche and Hargrove, 1999) and the body size and

survival of offspring (Jackson, 1952; Phelps and Clarke, 1974; Dransfield et al., 1989). These

seasonal changes in the fat body content may suggest seasonal differences in the tsetse’s

susceptibility to trypanosomal infections. Interestingly, a longitudinal study conducted on the

plateau of eastern Zambia showed a negative correlation between the fat body content and the

T. congolense infection rate of the G. m. morsitans population (Van den Bossche et al.,

unpublished work) (Figure 8.3) with the highest infection rates during the hot dry season

when tsetse’s survival is lowest.

Month

1 2 3 4 5 6 7 8 9 101112 1 2 3 4 5 6 7 8 9 101112 1 2 3 4 5 6 7 8 9 101112

Pro

port

ion

of

infe

cte

d G

. m

. m

ors

itan

s (

%)

0

2

4

6

8

10

Month

ly a

ve

rage f

at le

vel (m

g)

1,6

1,8

2,0

2,2

2,4

2,6

2,8

3,0

3,2Infection rate

Fat level

1991 1992 1993

Avera

ge a

mbie

nt

tem

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°C)

16

18

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24

26

28

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Figure 8.3: Monthly average fat body levels and T. congolense infection rates in G. m. morsitans

in the Eastern Zambia (Van den Bossche et al., unpublished)


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8.3.1.4. Climate change

Climate change is one of the components of the complex environmental changes occurring

around the world as a result of human activities. It is estimated that average global

temperatures will have risen by 1.0–3.5 °C by 2100 (Watson et al., 1996). In tropical Africa,

the equatorial countries such as Cameroon, Kenya and Uganda could experience rises of 1.4

°C by 2050 (Watson et al., 1996). For tsetse flies, the possible effects of climate change on

vectorial capacity are not known and need further investigations. However, as mentioned

above, climate change and the resulting increase in ambient temperature suggest a higher

metabolism and a higher susceptibility to trypanosomal infections. Such an increase in

susceptibility has been proven for a number of vector-borne diseases (Githeko et al., 2000).

8.3.2. Availability of hosts

For their survival, haematophagous tsetse flies require the regular presence of suitable hosts in

their habitat. This may be not much of a problem in extensive undisturbed areas where game

animal range freely. However, large areas in which tsetse flies occur are currently under

strong pressure from human activity with a concomitant rapid change in the environment.

Indeed, in large parts of tsetse-infected sub-Saharan Africa, the human encroachment and the

progressive clearing of the natural vegetation for cultivation has resulted in disappearance of

game animals (the natural hosts of tsetse) and the introduction of livestock. The resulting

habitat fragmentation may also reduce the availability of hosts as a result of the fly’s limited

capacity to move in a fragmented environment. For example, it has been shown recently that

the destruction and fragmentation of the natural habitat of tsetse due to the extensive clearing

of natural vegetation for cotton production on the plateau of eastern Zambia has resulted in a

significant reduction in the apparent density of tsetse compared to the areas where human

density is much lower and the natural vegetation largely undisturbed (Ducheyne et al., 2009).

In these highly fragmented areas, the residual tsetse population may suffer from a reduced

availability of hosts and nutritional stress with possible subsequent repercussions for their

susceptibility to trypanosomal infections. Although it is still premature, the high infection

rates in cattle kept in these highly fragmented areas supporting low densities of tsetse flies

suggests high infection rates of the flies and perhaps an increased susceptibility to infection.


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8.4. References

Aksoy, S., Gibson, W.C., Lehane, M.J., 2003. Interactions between tsetse trypanosomes with implications for the control of trypanosomiasis. Advance in Parasitology 53: 1-83.

Dransfield, R. D., Brightwell, R., Kiilu J., Chaudhury, M. F. B., Adabie, D. A., 1989. Size and mortality rates of Glossina pallidipes in the semi-arid zone of southwestern Kenya. Medical and Veterinary Entomology 3: 83-95.

Ducheyne, E., Mweempwa, C., De Pus, C., Vernieuwe, H., De Deken, R., Hendrickx, G., Van den Bossche, P., 2009. The impact of habitat fragmentation on tsetse abundance on the plateau of eastern Zambia. Preventive Veterinary Medicine 91: 11-18


Society for Tropical Medicine and Hygiene 55: 383-397.

Githeko, A.K., Lindsay, S.W., Confalonieri, U.E., Patz, J.A., 2000. Climate change and vector-borne diseases: a regional analysis. Bulletin of the World Health Organization 78: 1136-1147

Glasgow, J. P., Bursell, E., 1960. Seasonal variations in the fat content and size of Glossina

swynnertoni Austen. Bulletin of Entomological Research 51: 705-713.

Jackson, C.H.N., 1952. Seasonal variations in the mean size of tsetse flies. Bulletin of


Phelps, R.J., Clarke, G.P.Y., 1974. Seasonal elimination of some size classes in male Glossina morsitans morsitans Westw. Bulletin of Entomological Research 64: 313-324

Randolph, S.E., Williams, B.G., Rogers, D.J., Connor, H., 1992. Modelling the effect of feeding-related mortality on the feeding strategy of tsetse (Diptera: Glossinidae). Medical and Veterinary Entomology 6: 231-240

Rogers, D.J., Randolph, S.E., 1978. Metabolic strategies of male and female tsetse in the field. Bulletin of Entomological Research 68: 283-297

Van den Bossche, P., Hargrove, J.W., 1999. Seasonal variation in nutritional levels of male tsetse flies Glossina morsitans morsitans Westwood (Diptera: Glossinidae) caught using fly-rounds and electric screens. Bulletin of Entomological Research 89: 381-387

Watson, R.T., Zinyowera, M.C., Moss, R.H., 1996. Climate change 1995: impacts, adaptations and mitigation of climate change: scientific-technical analysis. Contribution of Working Group II to the second assessment report of the Intergovernmental Panel on Climate Change. Cambridge University Press, Cambridge, United Kingdon

Summary

Summary

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The obligate blood feeding tsetse fly (Diptera: Glossinidae) is an essential component in the

cyclical transmission of several African trypanosome species such as Trypanosoma

congolense and T. brucei spp., causing devastating diseases in both human and animal in sub-

Saharan Africa. The proportion of trypanosome-infected tsetse is a major determinant of the

transmission dynamics of the disease and is affected by various endogenous and exogenous

factors. Recently, the nutritional state of the flies at the time of their infective bloodmeal was

demonstrated to affect the susceptibility of teneral and older flies to infections with T.

congolense or T. b. brucei. The experimental work that is presented in this thesis aimed i) to

determine whether nutritional stress affects the ability of tsetse flies to develop human

pathogenic trypanosome species, ii) to explore the underlying mechanism that induce an

increased susceptibility in starved tsetse flies with focus on the tsetse fly’s immunological

abilities and finally iii) to determine the effects of seasonal climatic changes on the nutritional

status and immune peptide expression in field-caught tsetse flies in order to determine

whether changes in environmental conditions affect the trypanosome transmission dynamics

in the field.

The first chapter of the thesis reviews our current knowledge on the endogenous and

exogenous factors that affects tsetse-trypanosome interactions with special attention to tsetse-

related factors that interfere with the parasite midgut establishment and maturation in the

insect vector.

In chapter 2, starved and non-starved teneral and older flies were experimentally

infected to determine the effect of starvation on their susceptibility to T. b. gambiense or T. b.

rhodesiense. Use was made of tsetse species belonging to the savannah (G. m. morsitans) or

the riverine subgenera (G. p. gambiensis) in order to mimic the field situation as much as

possible. The results of this study revealed that starvation occurring before the infective

bloodmeal increases the intrinsic vectorial capacity of tsetse flies for T. b. rhodesiense by

lowering the developmental barrier especially at midgut level. Several experimental infections

using different T. b. gambiense strains were unsuccessful due to a very low ability of the

parasites to establish in the tsetse fly’s midgut. This demonstrates the difficulty in obtaining a

suitable T. b. gambiense-Glossina experimental model that allows the study of factors

influencing this parasite’s development.

In chapter 3, a study determining the effect of starvation of tsetse flies on the

maturation of a T. brucei brucei procyclic midgut infection into a metacyclic salivary gland

Summary

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infection is presented. Tsetse flies, infected as teneral, were starved for seven consecutive

days 10 days after the infective bloodmeal. Results of this experiment showed a significantly

increased proportion of flies with salivary gland infection in the nutritionally-stressed fly

group suggesting an enhanced maturation T. b. brucei infection due to the nutritional stress

during the critical period of salivary gland colonisation.

In the fourth chapter, the effect of nutritional stress experienced by adult reproducing

female tsetse flies on their offspring’ susceptibility to trypanosome infection is determined.

The results of this study revealed a significant increase in the intrinsic vectorial capacity, for

infections with T. congolense or T. b. brucei, of teneral flies emerging from starved adult

females compared to those emerging from non-starved females. This suggests that in the field

situation, environmental conditions that nutritionally stress adult reproducing female tsetse

flies may not only affect their reproductive performance but also increase the number of

infected flies that emerged from pupae produced by these stressed females.

Chapter 5 investigates the effect of nutritional stress on i) the non-induced baseline

gene expression level of the antimicrobial peptides attacin, defensin and cecropin in the tsetse

fly and ii) their induced expression level in response to bacterial (E. coli) or trypanosomal

challenge. Results show that starvation of newly hatched, unfed tsetse flies significantly

lowers their baseline antimicrobial peptide expression level especially for attacin and

cecropin. In response to trypanosome challenge, only non-starved older flies showed a

significant increase of the antimicrobial peptide gene expression within 5 days after ingestion

of a bloodmeal containing trypanosomes, especially so for T. brucei bloodstream forms.

These data suggest that a decreased immune gene expression level in newly hatched flies or a

lack of immune responsiveness of older flies to trypanosomes, as a result of fly starvation,

could be one of the contributing factors to the increased susceptibility of nutritionally-stressed

tsetse flies to a trypanosome infection.

Chapter 6 presents a study conducted in the Zambezi Valley (Zimbabwe) during the

hot dry season (corresponding to a period of increased nutritional stress for tsetse flies) and

the rainy season (corresponding to a period of lower nutritional stress for tsetse flies). The

study aimed at determining seasonal changes in the nutritional and immune status of tsetse

flies with a possible impact on the trypanosome transmission dynamic in the field. The results

show that the nutritional state and the expression levels of antimicrobial peptides attacin,

defensin and cecropin are reduced during the hot dry season compared to the rainy season,

especially in G. m. morsitans. However, the possible repercussions of such variation on

Summary

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trypanosome transmission by this tsetse species were difficult to assess due to the limited

number of tsetse caught and dissected which did not allow the establishment of an age-

infection prevalence relationship that would highlight changes in the susceptibility of

especially adult flies.

In the chapter 7, we investigated the effect of the developmental stage of a

monomorphic T. congolense strain in the mammalian host on its transmissibility by the tsetse

fly. The results showed that the development stage of the trypanosome in the host blood does

affect the proportion of flies that develop a mature or immature infection with significantly

higher infection rates in flies that were fed on days 5 or 10 post-infection of the mammalian

host (mouse). These findings stress the importance of standardising experiments in which the

vectorial capacity of tsetse flies is determined and compared.

In the final general discussion (chapter 8), the major findings of the thesis are

discussed in the wider context of the possible impact of nutritional stress and environmental

stressors on the epidemiological situation of sleeping sickness.

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110

De verplicht bloedzuigende tseetseevlieg (Diptera: Glossinidae) is een essentiële schakel in de

cyclische overdracht van verschillende Afrikaanse trypanosoomsoorten, waaronder

Trypanosoma congolense en T. brucei spp., die in het sub-sahara Afrika ernstige ziekten

veroorzaken bij mens en dier. De proportie van trypanosoom-geïnfecteerde vliegen in de

natuurlijke tseetseepopulatie is hierbij bepalend voor de dynamiek van de ziekte-overdracht

en wordt beïnvloed door verschillende endogene en exogene factoren. Recent werd

aangetoond dat de nutritionele conditie van de tseetseevlieg op het tijdstip van de infectieuze

bloedmaaltijd een belangrijke invloed heeft op de gevoeligheid van tenerale en oudere vliegen

om een infectie met T. congolense of T. brucei brucei te ontwikkelen. De experimentele

studies die in dit doctoraatswerk worden gepresenteerd hadden als uitgangspunt om i) na te

gaan of nutritionele stress de gevoeligheid beïnvloedt van de tseetseevlieg voor de

menspathogene Trypanosoma soorten, ii) het onderliggende mechanisme van de verhoogde

gevoeligheid bij nutritioneel gestresseerde vliegen te exploreren met een focus op de

immuunreactie van de tseetseevlieg en iii) na te gaan of seizoensgebonden

klimaatsveranderingen een impact hebben op de nutritionele conditie, immuunpeptide

expressie en trypanosoominfectiegraad van een natuurlijke tseetseevliegpopulatie. Dit laatste

zou ons mogelijk toelaten om in te schatten in hoeverre de dynamiek van de

trypanosoomoverdracht in het veld beïnvloed wordt door natuurlijke

omgevingsveranderingen.

In het eerste hoofdstuk geven we een literatuuroverzicht van onze huidige kennis in

verband met endogene en exogene factoren die een rol spelen in de tseetsee-trypanosoom

interactie. Extra aandacht wordt hierbij besteed aan tseetsee-gerelateerde factoren die de

ontwikkeling van de parasiet in de middendarm en de verdere maturatie beïnvloeden.

Het tweede hoofdstuk beschrijft experimenteel werk om het effect van nutritionele

stress op de ontwikkeling van de menspathogene parasieten T.b.gambiense en T.b.rhodesiense

in de vlieg na te gaan. Voor deze studies hebben we gebruik gemaakt van zowel Glossina

morsitans morsitans vliegen (savanah habitat) als van een ‘riveriene’ soort (G. palpalis

gambiensis) om de natuurlijke trypanosoom-tseetsee relatie beter te benaderen. Uit dit

experimenteel werk kwam naar voor dat nutritionele stress die de infectieuze bloedmaaltijd

voorafgaat, de intrinsieke vectoriële capaciteit van de tseetseevlieg verhoogt voor de T. b.

rhodesiense parasiet en dit als gevolg van een verlaging van de ontwikkelingsbarrière in de

middendarm. De infectie-experimenten met verschillende T.b.gambiense stammen waren

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111

echter niet succesvol door de zeer lage capaciteit van deze parasieten om zich in de

tseetseemiddendarm te vestigen. Dit laatste illustreert de moeilijkheid om een geschikt T. b.

gambiense-Glossina experimenteel model te identificeren dat de studie van factoren die de

parasietontwikkeling in de tseetseevlieg beïnvloeden mogelijk kan maken.

Het effect van nutritionele stress op de maturatie van een T. brucei brucei procyclische

middendarm infectie wordt beschreven in hoofdstuk drie. Tseetseevliegen, geïnfecteerd als

tenerale vlieg en die zeven dagen verhongerd werden 10 dagen na de infectieuze

bloedmaaltijd, ontwikkelden significant meer mature, metacyclische infecties in de

speekselklieren in vergelijking met de controle groep. Dit suggereert dat nutritionele stress

tijdens de kritische periode van de kolonisatie van de trypanosoom naar de speekselklieren

een bevorderende impact heeft op het maturatieproces van de parasiet in de tseetseevlieg.

In het vierde hoofdstuk beschrijven we het effect van nutritionele stress bij

reproductieve wijfjes op de gevoeligheid van hun nakomelingen om een trypanosoominfectie

te ontwikkelen. Uit deze studie komt duidelijk naar voor dat nakomelingen van nutritioneel

gestresseerde wijfjes significant meer infecties met T. congolense of T.b.brucei ontwikkelden

in vergelijking met nakomelingen van niet-gestresseerde wijfjes. Voor de veldsituatie

suggereert dit dat omgevingscondities die reproducerende wijfjes nutritioneel stresseren niet

alleen een directe impact zullen hebben op hun reproductieve capaciteit maar ook op de

proportie van vliegen (de nakomelingen) die zich met de trypanosoom parasiet zullen

infecteren.

Hoofdstuk vijf rapporteert het effect van nutritionele stress op i) het niet-geïnduceerde

basisniveau van genexpressie van de antimicrobiële peptiden attacine, defensine en cecropine

in de tseetseevlieg en ii) de geïnduceerde expressieniveaus in antwoord op een bacteriële

(E.coli) of trypanosoom blootstelling. De resultaten toonden ons dat nutritionele stress bij

nieuw uitgekomen, ongevoede vliegen de basis expressieniveaus significant verlaagt vooral

voor attacine en cecropine. Bovendien bleken enkel de oudere, niet gestresseerde vliegen een

significante immuunrespons vertonen vijf dagen na opname van een infectieuze

bloedmaaltijd, vooral voor de T.b.brucei parasiet. Deze data suggereren dat een lager

immuunpeptide expressie niveau bij pas uitgekomen vliegen of een gebrek aan immuunrepons

tegen de trypanosoom bij oudere vliegen, als resultaat van nutritionele stress, factoren zijn die

kunnen bijdragen tot de verhoogde gevoeligheid van nutritioneel gestresseerde tseetseevliegen

om een infectie met trypanosomen te ontwikkelen.

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Om na te gaan of de nutritionele conditie en de immuunstatus in een natuurlijke

tseetseepopulatie wordt beïnvloed door seizoensgebonden veranderingen werd er een

veldstudie uitgevoerd. De resultaten hiervan worden beschreven in hoofdstuk zes. Deze studie

werd uitgevoerd in de Zambezi vallei (Zimbabwe) gedurende het ‘hete-droge’ seizoen

(overeenkomend met een periode van verhoogde nutritionele stress voor de tseetseevliegen)

en het ‘regen’ seizoen (overeenkomend met een periode van lagere nutritionele stress). De

resultaten tonen dat de nutritionele conditie en de expressie niveaus van de antimicrobiële

peptiden attacine, defensine en cecropine verminderd zijn in het ‘hete-droge’ seizoen en dit

vooral bij de G. m. morsitans vliegen. Echter, de impact van deze veranderingen op de

dynamiek van de locale trypanosoomoverdracht door deze G. morsitans populatie blijft zeer

moeilijk in te schatten daar het aantal gevangen en gedissecteerde vliegen te beperkt was om

een accurate leeftijd- trypanosoom prevalentie relatie te kunnen opstellen.

Het zevende hoofdstuk beschrijft een studie in verband met de verdere optimalisatie

van het T. congolense-Glossina experimenteel model. Het effect van het

ontwikkelingsstadium van een monomorfe T. congolense stam in de zoogdiergastheer op de

overdraagbaarheid van de parasiet in de tseetseevlieg werd nagegaan. Hierbij toonden we aan

dat er significant meer tseetseevliegen een trypanosoominfectie ontwikkelen wanneer ze hun

infectieuze bloedmaaltijd nemen op dag 5 of dag 10 van de infectie bij de zoogdiergastheer

(muis). Deze resultaten benadrukken nog maar eens het belang van de standaardisatie van de

infectie experimenten die we uitvoeren om de vectorïele capaciteit van tseetseepopulaties te

bepalen en te vergelijken.

In het laatste hoofdstuk geven we een synthetiserende discussie waarbij we de

belangrijkste bevindingen van ons experimenteel werk kaderen in een bredere context van de

mogelijke impact van nutritionele stress en omgevingsstressoren op de epidemiologische

situatie van slaapziekte.

Nutritional stress affects the tsetse fly's immune gene expression

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