hytrosavirus
2009, he obtained an MSc degree in Cellular/Molecular Biotechnology (Wageningen University, The Neth-erlands), sponsored by The WU MSc Scholarship. In September 2010, he started his PhD at the Laboratory of Virology, Wageningen University, on a WU PhD Fellowship, and in March 2011, he transferred to NUFFIC PhD Glossina hytrosavirus and explore methods to control the virus in tsetse massproduction facility at Seiber -sdorf. His research at Wageningen University’s Virology and IAEA’s Seibersdorf Laboratories resulted to this thesis. Henry also spent part of his PhD study time as1. Hytrosaviruses cannot be used as bio-control agents for tsetse flies. (this thesis).
2. As an insect host metamorphoses to adulthood, vertical transmission of an insect virus becomes epizootically more important than horizontal transmission.
(this thesis).
3. Microbial symbiosis is a remarkable source of evolutionary innovation.
4. The contents of a proteome and the repertoire of domains in the protein sequences can be used to trace the evolutionary history of an organism.
5. The presence of protein-coding regions or open reading frames in a genome does not necessarily imply the presence of functional proteins.
6. Understanding the link between a rewarding social interaction and a lasting behavioural change may provide a solution to depression and drug abuse in humans.
(Shohat–Ophir et al., (2012), Science 335: 1351–54).
7. The impression that hard work is degrading to fashionable life does not appeal to reason.
8. The maxim that a slave first loses his name and then adopts the lingo is true in all facets of life. (Inference from Mũrogi wa kagogo, a Kikuyu satire by Ngũgĩ wa Thiong'o, 2006).
Proposition belonging to the thesis
Glossina hytrosavirus control strategies in tsetse fly factories:
Application of infectomics in virus management
Henry Muriuki KARIITHI
Glossina
hytrosavirus
control
strategies
in
tsetse
fly
factories:
Application
of
infectomics
in
virus
management
Thesis committee
Promoters Prof. dr. J.M. Vlak
Personal Chair at the Laboratory of Virology Wageningen University
Prof. dr. M.M. van Oers Professor of Virology Wageningen University
Thesisco‐promoters
Assoc. Prof. dr. Adly M.M. Abd‐Alla )nsect Pest Control Laboratory
)nternational Atomic Energy Agency, Austria
Dr. Grace Murilla Centre Director
Trypanosomiasis Research )nstitute
Kenya Agricultural Research )nstitute, Kenya
OtherMembers
Prof. dr. W. Takken, Wageningen University
Prof. dr. D. G. Boucias, University of Florida, Gainesville, USA
Prof. dr. J. Van den Abbeele, )nstitute of Tropical Medicine, Antwerpen, Belgium Prof. dr. S.C. de Vries, Wageningen University
Glossina
hytrosavirus
control
strategies
in
tsetse
fly
factories:
Application
of
infectomics
in
virus
management
Henry
Muriuki
Kariithi
Thesis
submitted in partial fulfilment of the requirements for the degree of doctor at Wageningen University
by the authority of the Rector Magnificus Prof. dr. M.J. Kropff
in the presence of the
Thesis Committee appointed by the Doctorate Board to be defended in public
(. M. Kariithi
Glossinahytrosavirus control strategies in tsetse fly factories:
Application of infectomics in virus management
pages
Thesis, Wageningen University, Wageningen, NL
With references, with summaries in English, Dutch and Swahili.
Abstract
African trypanosomosis is a fatal zoonotic disease transmitted by tsetse flies Diptera; Glossinidae , blood‐sucking insects found only in sub‐Sahara African countries. Two forms of trypanosomoses occur: the animal African trypanosomosis AAT; nagana , and the human African trypanosomosis (AT; sleeping sickness . Since there are no effective vaccines against trypanosomosis, tsetse eradication is the most effective disease control method. Tsetse can be effectively eradicated by the sterile insect technique S)T , which is applied in an area‐wide integrated pest management approach. S)T is an environmentally benign method with a long and solid record of accomplishment. S)T requires large‐scale production of sexually sterilized male flies usually by exposure to a precise and specific dose of ionizing radiation, usually from a
Co or Ce source , which are sequentially released into a target wild insect
population to out‐compete wild type males in inseminating wild virgin females. Once inseminated by sterile males, the virgin females do not produce viable progeny flies. )mportantly, these females do not typically re‐mate. Ultimately, the target wild insect population can decrease to extinction. (owever, tsetse S)T programs are faced with a unique problem: laboratory colonies of many tsetse species are infected by the
Glossinapallidipessalivary gland hypertrophy virus GpSG(V; family Hytrosaviridae .
GpSG(V‐infected flies have male aspermia or oligospermia, underdeveloped female ovarioles, sterility, salivary gland hypertrophy syndrome SG( , distorted sex ratios, and reduced insemination rates. Without proper management, SG(V can cause
collapse of Glossina pallidipes colonies. To ensure colony productivity and survival,
GpSG(V management strategies are required. This will ensure a sustained supply of sterile males for S)T programs. The aim of this PhD research was to investigate the functional and structural genomics and proteomics infectomics of GpSG(V as a prerequisite to development of rationally designed viral control strategies: A series of experiments were designed to: i investigate epidemiology and diversity of GpSG(V; ii identify GpSG(V proteome and how viral and host proteins contribute to the pathobiology of the virus; and iii investigate the interplay between GpSG(V, the microbiome and the host, and how these interactions influence the outcomes of viral infections. By relating GpSG(V and host infectomics data, cost‐effective viral management strategies were developed. This resulted in significant reduction of
Table
of
Contents
Abstract
Chapter1 General )ntroduction
Chapter2 Dynamics of GpSG(V transmission in G.pallidipes colonies
Chapter3 Prevalence and diversity of GpSG(V in wild G.pallidipes
populations
Chapter4 Proteome and virion components of GpSG(V
Chapter5 Salivary secretome of GpSG(V‐infected G.pallidipes
Chapter6 Role of microbiome in GpSG(V transmission
Chapter7 Management of GpSG(V infections in G.pallidipescolonies
Chapter8 General Discussion
Chapter
1
General
introduction
Introduction
This PhD dissertation covers various aspects of the pathobiology, epidemiology,
morphology and morphogenesis of the Glossinapallidipessalivary gland hypertrophy
virus GpSG(V , a double‐stranded DNA dsDNA virus classified in the Hytrosaviridae
family of insect viruses )CTV; Abd‐Alla etal., b . GpSG(V is a major pathogen of
laboratory colonies of the tsetse fly G.pallidipes Diptera; Glossinidae . Typically, a
small proportion of laboratory G.pallidipes flies infected by GpSG(V develop
hypertrophied salivary glands and midgut epithelial cells, and show gonadal/ovarian anomalies, distorted sex‐ratios, reduced insemination rates, fecundity and lifespan. These symptoms are rarely observed in wild tsetse populations.
)n East Africa, G.pallidipes is one of the most important vectors of the debilitating
zoonotic disease, African trypanosomosis. A large arsenal of tsetse and trypanosomosis management tactics is available. The sterile insect technique S)T is a robust control and effective method to eradicate tsetse fly populations when integrated with other control tactics in an area‐wide integrated approach. S)T requires production of sterile male flies in large‐scale production facilities. To supply sufficient
numbers of sterile males for the S)T component against G.pallidipes, strategies have to
be developed to manage GpSG(V infections in the fly colonies. This chapter provides a historic account of tsetse fly and trypanosomoses control, and a chronology of the emergence and biogeography of hytrosaviruses. The thesis rationale is introduced.
African
trypanosomoses
–
the
"neglected
tropical
diseases"
Tsetse flies are important vectors of two debilitating diseases; human African trypanosomosis (AT or sleeping sickness , and African animal trypanosomosis AAT
or nagana Mattioli etal., . Tsetse flies and trypanosomosis render vast areas of
agricultural land un‐exploitable, especially during the rainy seasons Mamoudou etal.,
. Although there are over species and sub‐species of tsetse, most of which can transmit trypanosomosis, only ‐ of these species are of medical and agricultural
importance. The most important tsetse vectors are the riverine species G.palpalis,
G.morsitans, G.austeni and G.pallidipes in Eastern and Southern Africa Smith etal., . Although tsetse fly fossils have been found in ‐million‐year‐old shales of
Florissant, Colorado, USA Cockerell, , to date, tsetse flies are confined to Africa
except from an isolated population on the Arabian Peninsula Elsen etal., .
(AT is one of the most serious of the so‐called 'neglected tropical diseases' NTDs
(otez and Kamath, . NTDs are a group of chronic diseases endemic in low‐
income populations in Africa, Asia and the Americas (otez et al., .
Trypanosomosis is restricted to sub‐Saharan African countries and its distribution
extends to more than million square kilometres of the African continent Cecchi et
al., Figure1 .
The people at the highest risk of tsetse bites and of contracting (AT are the rural populations that primarily depend on small‐scale agriculture, fishing, animal husbandry and hunting. Resurgence and epidemics of (AT are often associated with economic decline, civil disturbances/wars, population movements and refugees
Smith etal., . The presence of tsetse and trypanosomosis is considered as one of
Figure 1: Distribution of differenttsetseflyspecies in sub‐Saharan African countries. The different colour in the figure legend represents the different tsetse fly species in the map
the "roots of hunger and poverty" in sub‐Saharan Africa Vreysen, . )t is estimated that approximately % of Africa s livestock consists of herds in small
villages Otte and Chilonda, . This implies that maintaining healthy animals can
make the difference between subsistence misery and an acceptable lifestyle for the farmers and their families. FAO estimates that ~ US$ . billion worth of agricultural products are lost annually due to AAT including ~ million cattle deaths , and human lives are lost daily due to (AT.
(AT is difficult to treat, and there are no effective vaccines available against both AAT and (AT. None of the available trypanocidal drugs for (AT is ideal; their treatment schedules are prolonged, excruciatingly painful often described by patients as "fire in
the veins" and require continuous hospitalization Matovu etal., . Generally, the
tsetse‐transmitted trypanosomes initiate their lifecycle by first colonizing the tsetse hosts midguts, and then migrate into the ectoperitrophic space, and to the salivary
glands via the alimentary canal and the mouth parts Oberholzer etal., . The
parasites differentiate into the final mammalian‐infective trypomastigocyte in the tsetse salivary glands, and are then transmitted to the mammalian host by an infected
tsetse bite Sharma etal., . )t should however, be noted that some steps in the
lifecycle of trypanosomes are group‐specific. For instance, members of the T.vivax
group only stay in the proboscis; the T.congolensegroup has a cycle involving the
proboscis and the midguts; while only the T.bruceigroup has a cycle involving the
midgut and the salivary glands Vickerman, .
Without treatment, (AT can be fatal. (owever, fatalities of trypanosomes differ from
one group to another. For instance, in West Africa T.b.gambiensis causes a chronic
(AT that can take many years to kill a patient, while in East Africa, T.b.rhodiensis
causes an acute (AT that can kill a patient within weeks Brun etal., ; Chappuis
etal., . The most widely used drug, melarsoprol, which was developed in
Friedheim, , is lethal for up to % of the treated patients Burri etal., ;
. )t is also important to note that, one of the biggest problems in the treatment of (AT is that the patients are usually so weak that they are more likely to die from the treatment rather than from the disease. )n addition, patients need to be properly fed for several weeks to regain strength before the commencement of treatments. This presents a very serious problem considering that there are hardly any available funds to properly feed, or purchase drugs for (AT treatments. Besides, the available drugs for AAT are overly expensive for African peasant farmers, and there are reports of
increasing drug‐resistance and drug‐counterfeiting Barrett etal., ; Geerts etal.,
. For all the above‐mentioned reasons, control of the disease vector tsetse is of critical importance, and probably represents the most sustainable trypanosomosis
An
overview
of
tsetse
fly
control
methods
Two main characteristics of the tsetse fly render them suitable for eradication. Firstly, compared to other insects of medical and agricultural importance, tsetse flies have
low dispersal and reproduction rate k‐strategists Leak, : tsetse flies are
viviparous bearing live young , and typically produce ‐ offspring in their lifespan
Attardo et al., . Therefore, unlike many insect vectors that produce large
numbers of eggs r‐strategist , tsetse flies have limited capacity to rebound in areas
where their populations have been reduced. Secondly, tsetse flies are adapted for efficient exploitation of stable habitats provided by vertebrate nests or human dwellings with low levels of cross‐breeding. This means that tsetse flies have reduced genetic variability within each vector population and therefore, have limited capacity to respond through selection pressure to various control interventions Dujardin and
Schofield, .
Tsetse control methods have evolved from discriminate bush clearing and wild game
culling at the beginning of the th Century, to the widespread applications of broad‐
spectrum insecticides after the Second World War Allsopp, . (owever, these
control measures have negative impacts on the environment and ecosystems, and are not compatible anymore with today s environmental requirements. The overall effect of bush clearing and game culling is direct loss of biodiversity. The use of insecticides
raises concerns on the fate of non‐target organisms du Toit, . Besides, some
insecticides persist long in the environment, and the residues of some of these insecticides end up in water bodies where they endanger aquatic organisms, or are
transferred and concentrated up the food chains Grant, . To respond to the
above‐mentioned negative environmental and ecological impacts, advancements were made in using traps, insecticide‐impregnated targets Brightwell and Dransfield,
and live bait technologies Rowlands etal., . Although these methods have
been used successfully to reduce local tsetse populations Leak, , each of these
methods has its own limitations. Firstly, the methods do not protect the cleared areas from re‐invasion by tsetse flies from residual pockets and from neighbouring
territories Brightwell et al., . Secondly, the methods are applied in
administratively‐defined regions and run for an administratively‐specified time
Schofield and Kabayo, , which mostly depend on how long external donor funds
Application
of
the
sterility
principle
for
tsetse
eradication
)n , Knipling developed the theory of controlling insect pest population by
manipulating their reproductive capacity. (e likewise modelled that a target population could be eradicated when the release of sterile males was applied on an area‐wide basis against an entire insect pest population in a delineated area Knipling,
; , rendering them sterile. This method, commonly known as the sterile
insect technique S)T , involves large‐scale production of insects in mass‐rearing facilities. Excess male flies are sexually‐sterilized by exposure to a precise and specific
dose of ionizing radiation, usually from a Co or Ce source Robinson, ; .
The sterile males are then sequentially released into the target insect population in
numbers that allows them to out‐compete wild males for wild virgin females Abila et
al., . After the virgin females have mated with the sterile males, embryogenesis
is arrested, and consequently no viable offspring is produced. When the release of the sterile males is sustained, the size of the target insect population will decline and can eventually become extinct.
The S)T is a robust control tactic that has been used very successfully against insect pests that are important for agriculture and trade. As an example, the S)T was used to
control the screwworm fly Cochliomyiahominivorax Diptera; Cochliomyia from the
Southern USA, Mexico, Central America and Panama Wyss, and from northern
Libya after a serious outbreak in Lindquist etal., . The S)T was also used
to eradicate or suppress Mediterranean fruit fly Ceratitiscapitata Diptera; Ceratitis
populations in Chile, Mendoza Argentina , Mexico etc., and in Central America, South
Africa, )srael etc., respectively Enkerlin, ; Franz, . Lately, the S)T has also
been used with great success against several Lepidopteran pests such as the codling
moth Cydia pomonella L. in the Okanagan Valley of Canada Bloem et al., a;
b , the false codling moth Thaumatotibialeucotreta Meyrick in South Africa, the
Australian painted apple moth Teiaanartoides Walker in New Zealand Vreysen etal.,
, and the pink bollworm Pectinophora gossypiella Saunders in Texas, New
Mexico, Arizona, California USA and in Sonora and Chihuahua of northern Mexico
Enkerlin, ; Koyama etal., .
The S)T has also played a pivotal role in the sustainable area‐wide eradication of the
tsetse fly Glossinaausteni from Unguja )sland Zanzibar Vreysen etal., . This
program was preceded by successful applications of the technique against
G.palpalisgambiensis andG.tachinoides in the Sideradougou area in Burkina Faso and
against G.palpalispalpalis in the Lafia area of Nigeria Oladunmade et al., ;
Politzar etal., . The programs in Burkina Faso, and Nigeria were however not
implemented according to AW‐)PM principles and the tsetse‐cleared area was re‐ invaded by tsetse flies after the programs were completed. Following the area‐wide
The success of S)T in eradicating G.austeniand trypanosomosis from Unguja )sland inspired African Governments to call for increased efforts to manage the tsetse fly and trypanosomosis on mainland Africa. Consequently, an AW‐)MP program with an S)T
component was initiated in to eradicate G.pallidipes from a , square
kilometres of under‐utilized fertile land in the Southern Rift Valley of Ethiopia. For the Ethiopian S)T program, laboratory tsetse colonies were established at )nsect Pest Control Laboratory )PCL of the Joint FAO/)AEA Agriculture and Biotechnology
Laboratories in Seibersdorf, Austria, and at Kality, Ethiopia Feldmann et al., .
(owever, efforts to establish mass‐rearing of tsetse colonies revealed that G.pallidipes
colonies are vulnerable to a virus infection that causes salivary gland hypertrophy syndrome SG( , and leads to reduction in productive fitness in male and female
tsetse flies Abd‐Alla et al., b . Although the virus infection does not cause fly
mortalities, it results in significant reduction in fertility and fecundity leading to
decline of the colonies within a few generations Kariithi etal., a .
Discovery
and
distribution
of
hytrosaviruses
Chronological developments from the first emergence of the SG( syndrome to the
discovery of the hytrosaviruses causing the syndrome are summarized in Table1.
The first description of SG( syndrome was reported in G.pallidipesby Whitnall in the
s Whitnall, ; during investigations of Trypanosoma‐infections in
Zululand, South Africa. The syndrome was later described to be sex‐linked, and to
favour development of trypanosomes in G.morsitans Burtt, . )n the s, the
SG( syndrome was associated with a virus found in cytoplasmic vacuoles of the
salivary glands and midgut epithelial cells of G.fuscipes and G.morsitans Jenni, ;
; a; b . The virus was at that time described as virus‐like particles
VLPs , morphologically resembling those that had been described in Drosophila,
Aedesaegypti, and in nematodes. Since other hematophagous insects such as mosquitoes, ticks, sand flies and gnats had been widely known to transmit arboviruses, the virus particles found in tsetse were erroneously suggested to be an arbovirus. Other features of the tsetse virus particles that led to this conclusion is the shape, size of the viral particles, and the fact that arboviruses were known to replicate
in the host salivary glands Chamberlain, ; Janzen etal., ; Mims etal., ,
followed by release of mature virus particles via saliva during feeding La Motte, . A resemblance of the tsetse virus to baculovirus was also suggested on the
basis of their extended rod‐shape morphology Jaenson, b . Mating experiments
revealed that the virus is transmitted from the mother to her progeny and not from
the male parent Jaenson, . )n exceptional cases, symptomatic SG( ‐progenies
were produced by asymptomatic parents, meaning that the virus is possibly reactivated from latency by a combination of stress and genetic factors. (igh
that SG( contributes to natural regulation of tsetse populations in the field Odindo, . The SG( syndrome was reported to occur twice as frequent in males than females, expressed in one‐third of the flies that had been fed or micro‐injected the virus suspension as tenerals newly‐ emerged and non‐fed flies , and to cause female
sterility, and male aspermia and/or oligospermia Odindo etal., .
Table1:Chronological history of the discovery and distribution of salivary gland hytrosaviruses SG(Vs .
Investigator(s) Year Majorcontribution(s) References
Whitnall , First published record of SG( Glossinaspp. Whitnall, ; Burtt , Suggested that SG( is sex‐linked Burtt, Jenni etal., , , Described virus particles in G.morsitans and
G.fuscipesfuscipes; suggested Golgi‐ER viral assembly Jenni, b ; ; a; Lyon First published record of SG( in M.equestris Lyon,
Jaenson First clear association of viral particles with SG( Jaenson, b Amargier etal., Reported SG( in M.equestris Amargier etal., Otieno etal., Reported SG( as common feature in wildG.pallidipes Otieno etal., Opiyo Reported poor productivity of G.pallidipescolony at
Kenya Trypanosomiasis Research )nstitute KETR) , Kenya
Opiyo and Okumu,
Odindo etal., , , Demonstrated that viral particles are infectious peros;
First report that Glossina virus has dsDNA genome Odindo
etal., ; ;
; Jaenson First report on reduced insemination rates, fecundity
and lifespan in laboratory colonies of G.pallidipes Jaenson,
Ellis etal., Reported SG( in Zimbabwe and )vory Coast Ellis and Maudlin, ; Gouteux,
)AEA , Reported poor productivity of G.pallidipescolonies at )nsect Pest Control Laboratories )PCL , Seibersdorf, Austria
Odindo Proposed Glossina virus as a bio‐control agent Odindo, Jura etal., , ,
, Demonstrated transmission of artificial infection Glossinavirus after Jura ; etal., ; ; Kokwaro etal., ‐ Cytopathology of virus particles in tsetse salivary glands Kokwaro etal., ; Shaw Reported SG( in G.m.Swyenatoni and G.brevipalpis Shaw and Moloo, Coler etal., First published record of SG( in M.domestica Coler etal., Sang etal., ‐ Reported SG(V in tsetse milk glands, mid‐gut and male
accessory reproductive glands Sang
etal., ; ;
; )AEA Collapse of an Ethiopian‐derived G.pallidipescolony at
)PCL, Seibersdorf, Austria
Kokwaro Reported virus particles in male accessory reproductive
glands of G.m.morsitansWestwood Kokwaro, Abd‐Alla etal.;
Garcia‐Maruniak
etal.,
G.pallidipesand M.domesticaSG(Vs genome sequenced Abd‐Alla etal., ; b
Abd‐Alla etal., Establishment Hytrosaviridaefamily Abd‐Alla etal., b Salem etal., Transcription analysis of M.domesticaSG(V Salem etal., Kariithi etal., ‐ Described the proteome, ultra‐structure and
morphogenesis of Glossinavirus Kariithi etal., b; Prompiboon et
al., Reported wild‐wide distribution of SG(V in M.domestica Prompiboon etal.,
Luo and Zheng SG(V‐like virus described in accessory gland filaments
of the parasitic wasp D.Longicuadata Luo and Zeng,
Boucias etal., Described the role of endosymbionts on
trans generational trans mission of SG(V in G.pallidipes Boucias
etal., b Abd‐Alla etal., Reported successful management of GpSG(V and
eradication of SG( in G.pallidipescolonies Abd‐Alla
)t was not until that the virus causing SG( syndrome in G.pallidipesAusten was discovered to be a novel DNA virus, which could not be placed in any of the existing
taxa of insect DNA‐viruses Odindo etal., . After purifying the virus by a series of
sucrose gradient centrifugations, the virus particles were described as long, non‐
enveloped rods containing linear double stranded ds DNA and different
polypeptides Odindo etal., .
The second description of SG( syndrome was reported in adult populations of the
narcissus bulb fly, Merodonequestris Diptera; Syrphidae in the s in southern
France Amargier etal., ; Lyon, ; Lyon and Sabatier, . The incidence of
SG( was reported in % and % of M.equestrisnobilis and M.equestristransversalis
Diptera; Syrphidae , respectively. Degeneration of male and female reproductive
organs Lyon, was also described as the main disease symptom. Similar to the
virus causing SG( in tsetse flies, the virus particles in M.equestrishad ultra‐structural
features similar to some baculoviruses, and were therefore, assumed to be related to tsetse hytrosavirus. To date, no further research has been performed on the bulb fly virus to substantiate this claim.
The third discovery of an SG( syndrome was reported in the s by Coler etal.,
Coler etal., in adult house flies, Muscadomestica Diptera; Muscidae , during a
survey of parasitic nematodes in the fly at a dairy farm in central Florida, USA. Similar to the tsetse fly, SG( syndrome and total suppression of oogenesis were described as
the main symptoms of housefly virus infection Coler etal., . (owever, unlike in
tsetse fly virus, there is no evidence of vertical transmission of the housefly virus from
mother to the progeny Geden etal., ; Lietze etal., . M.domesticawas later
confirmed to be naturally infected with the housefly virus, and the virus has been shown to be globally distributed with detections in samples from Africa, North
America, Europe, Asia, the Caribbean, and the south‐western Pacific Geden et al.,
b; Prompiboon etal., . )n addition, the housefly virus was also reported to
replicate in a laboratory colony of the stable fly, Stomoxyscalcitrans Diptera;
Muscidae , which occurs sympatrically with the house fly, albeit without the classical
SG( syndrome Geden etal., a .
Recently, a virus with similarity to those in the hytrosavirus group was accidentally
discovered in the accessory gland filaments of the braconid wasp Diachasmimorpha
longicuadata (ymenoptera; Braconidae , in a sample derived from a population
originally from (awaii, released in Thailand and introduced to China as a bio‐control
for the fruit fly, Bactrocelladorsalis Diptera; Tephritidae in South China Luo and
Zeng, . Although the virus was detectable in the accessory gland filaments
unlike the salivary glands in dipteran insects described above , the wasp accessory glands appeared hypertrophied with ultra‐structural features similar to
Pathology
of
hytrosaviruses
Dipteran adult flies infected by hytrosaviruses show gross signs of overt salivary gland hypertrophy symptom SG( , hence the name SG(Vs. Of the SG(Vs, the tsetse fly and housefly viruses have so far been the most widely studied. The two hytrosaviruses induce similar gross pathology in their hosts, most notably the characteristic hypertrophy of salivary glands of the adult insects and reduction in
reproductive fitness Abd‐Alla etal., b; Lietze etal., . Pathological effects of
SG(Vs have been observed more profoundly in laboratory‐bred tsetse fly colonies. )n
, wild‐caught G.pallidipesfrom the Kibwezi forest, Kenya, were used to initiate a
colony at the Kenya Trypanosomiasis Research )nstitute KETR) . Within two years of its establishment, the colony declined due to poor productivity Opiyo and Okumu,
. A similar trend was noted in another G.pallidipes colony established at the
)nsect Pest Control Laboratories )PCL in Seibersdorf, Austria, which experienced a
steady decline, eventually leading to its collapse in Abd‐Alla et al., a;
b . )nvestigations revealed that % of the males and % of the females had SG( syndrome. Tsetse with SG( syndrome exhibit discoloured bluish‐white salivary
glands that are enlarged than times larger than the normal thickness Figure2A .
Figure2:Thepathologyofhytrosaviruses. A Normal Nsg and hypertrophied (sg salivary glands dissected from G.pallidipes. )t should be noted that the pair of Nsg are dissected from a different fly for comparison with the (sg. Notice that the glands exhibiting SG( are enlarged times the size of normal glands; B Male G.pallidipes with asymptomatic i and symptomatic ii salivary glands. C Female
M.domesticawith healthy and D hypertrophied salivary glands showing lack of ovarian development in
Although there are no obvious external signs of infected flies Odindo, , hypertrophied glands appear as a pale outline in the male fly s abdomen with
irregular ridges on the cuticle Figure2B . The discolouration is probably due to the
extension of gland cells towards the cell lumina, resulting to constricted gland lumens. The enlarged and chalky‐white glands have also been observed in virus‐infected
house flies Figure2C and D Coler et al., . The collapse of G.pallidipes
colonies was ascribed to low productivity due to male testicular degeneration and
female ovarian abnormalities caused by the Glossina virus Jura etal., ; Kokwaro,
; Kokwaro and Murithi, ; Kokwaro and Odhiambo, ; Sang etal., ;
.
Glossina
hytrosavirus
as
tsetse
bio
‐
control
agent
Many insect‐pathogenic viruses such as baculoviruses are effective bio‐control
agents against insect pests (arrison and (oover,
; Moscardi,
.
Thepotential of the Glossina virus as a male sterility factor in tsetse control was first
proposed by Odindo . After micro‐injection of the virus into laboratory‐bred
G.pallidipes, infection was observed in . % and . % of treated male and female parents, respectively. The prevalence of SG( in the F progeny adults was much higher
than in the parents % in males and . % females . Whereas all infected females
were fertile, all infected males had SG( syndrome and were sterile. Maternal larviposition, F pupae weight and F pupae incubation periods were normal regardless of treatment. Two other studies reported that virus‐infected males showed
reduced reproductive potential Sang et al., . Based on these results, it was
hypothesized that the GlossinaSG(V may be applied as a tsetse bio‐control agent as
follows: the sterile male parents might compete with normal wild males in mating, and the fertile but infected females might transmit the SG( syndrome trans‐ovarially to subsequent generations, since such females produce only infected progeny
Jaenson, , where males are "born" sterile.
(owever, application of the Glossina SG(V as bio‐pesticide for tsetse control is
technically challenging. Firstly, recent findings show that neither micro‐injection nor
per os infection of the virus in G.pallidipes result in SG( syndrome in the same
parental generation, rather, the syndrome is only detectable in the third ~ %
and fourth ~ % larviposition cycles of the F generation Boucias etal., b .
Secondly, there is available evidence that, high prevalence of SG( in G.pallidipes
reduces the mating propensity and competitiveness of males thus affecting the
stability and performance of tsetse colonies Mutika et al., , and hence
difficulties in producing high amounts of infected insects. Thirdly, in vitro mass
to produce the virus in an alternative host house fly ‐ with short life cycle and easy
to produce enmasse ‐ have been unsuccessful. Fourthly, there is no available evidence
for horizontal transmission of GlossinaSG(V through contact between flies, mating, or
faecal contamination, thus limiting the modes of how the virus would be dispersed in the field. Finally, the virus does not produce occlusion bodies, as for instance baculoviruses do to achieve prolonged stability in the environment outside of the
host: recent evidence shows that GlossinaSG(V is highly unstable outside of the host
Kariithi et al., b , with more than % of purified virus suspension losing
infectivity after three days at °C Abd‐Alla et al., b . Formulation of virus
suspensions allowing virus to retain infectivity under both laboratory and field conditions, thus appears insurmountable for the time being. This is in contrast to the
Muscavirus infection system: intra‐hemocoelic injection with very low dosage of the
virus induces both % incidence of SG( syndrome, and a total shut down of
oogenesis within ‐ h post‐injection Geden etal., a; Lietze etal., ; Lietze
et al., . Therefore, the use of the Glossina SG(V as tsetse bio‐control agent
appears impractical.
Genome
organization
of
hytrosaviruses
The negative impacts of Glossina SG(V infections on laboratory‐bred G.pallidipes
prompted researches to understand the viral biology and pathology. (ypertrophied
salivary glands were dissected from G.pallidipes flies originating from Tororo, Uganda
in , colonized initially at Leiden University, The Netherlands, and subsequently
transferred to )PCL, Seibersdorf, Austria in . The genome of the virus purified
from the dissected glands was fully sequenced NC_ . Abd‐Alla etal., .
A total of non‐overlapping open reading frames ORFs were identified, of which
ORFs were presumed to encode putative viral proteins Abd‐Alla etal., .
Detection of two bands super coiled and relaxed forms after agarose gel electrophoresis of purified DNA, and a lack of end‐labelling of the undigested DNA
suggested a circular viral genome Figure3 . One hundred thirteen . % of these
ORFs did not match to any of the sequences available in various databases Abd‐Alla et
al., . Thirty‐seven ORFs . % were homologues to genes of other viruses,
while ten . % were homologues to non‐viral/cellular genes. Most notable of the
tsetse virus ORFs that were homologues to other viral genes were five of the peros
infectivity factor genes pifs p74, pif‐1, pif‐2, pif‐3, and odv‐e66 encoded by
baculoviruses and nudiviruses Song etal., . Other notable homologies included
homologues to; sixteen entomopoxvirus and poxvirus genes, three iridovirus and nimavirus genes each, two ascovirus genes and one herpesvirus gene. Most notable of
the cellular gene homologues include chitinase str. G.m.morsitans , DNA helicases,
thymidylate synthases, and several homologues to bacterial genes Abd‐Alla et al.,
sequence and fourteen direct repeat sequences drs composed of ‐ bp units.
Figure3:CircularrepresentationofGlossinaSGHVgenome. Arrows indicate position and direction of transcription for the potential ORFs. GpSG(V ORF numbers and putative genes are shown. The alphabetical numbers represent restriction fragments generated by Bg))) enzyme during the electrophoretic profiling of the virus genome.
The Glossinavirus genome was found to be a circular dsDNA molecule of , bp,
with the putative ORFs distributed equally on both strands % forward, %
reverse and a gene density of one ORF per . kb Abd‐Alla etal., . Position one
protein. Many of the ORFs are clustered into inferred cassettes in both strands, and
represent % of the genome. The genome has a high A+T content % . About %
of the viral genome is composed of repeat sequences in total , consisting
essentially of direct repeats, distributed throughout the genome.
As reported earlier Odindo etal., , when GlossinaSG(V was identified the virus
could not be assigned to any of the families of DNA viruses described at that time
Abd‐Alla etal., when considering its signature characteristics: induction of SG(
syndrome, possession of an enveloped rod‐shaped virus particle, a large circular
dsDNA genome, and being non‐occluded Abd‐Alla et al., a . Based on these
characteristics, the virus was proposed to be accommodated in a new virus family,
Hytrosaviridae, a name derived from "Hypertrophia sialoadenitis", the Greek word for
"salivary gland inflammation". The Glossina SG(V is commonly referred to as the
salivary gland hypertrophy virus GpSG(V , and is classified in the newly‐established
Hytrosaviridaefamily, genus Glossinavirus, and species Glossinahytrosavirus Abd‐Alla
etal., b . This taxonomy is now accepted by the )CTV http://ictvonline.org/ .
Phylogeny
of
hytrosaviruses
Phylogenetic analysis of (ytrosaviruses SG(Vs based on the DNA polymerase gene, which is present in all large dsDNA viruses, does not cluster these hytrosaviruses with
other insect dsDNA viruses Abd‐Alla et al., ; Garcia‐Maruniak et al., .
)nstead, the DNA polymerase of SG(Vs clusters more closely to that of herpesviruses and other viruses with linear dsDNA genomes. On the other hand, the alignment‐free method using whole proteome phylogenetic analyses of dsDNA viruses shows close association of the SG(Vs and nimaviruses specifically the white spot syndrome virus;
WSSV Gao and Luo, ; Wu et al., ; Yu et al., . Despite the apparent
ambiguities, these and other phylogenetic methods, such as super‐tree and super‐
matrix methods Wang and Jehle, ; Wang etal., , support the notion of a
common ancestry of SG(Vs with baculoviruses, nudiviruses and nimaviruses Jehle et
al., ; Wang et al., . As mentioned above, SG(Vs have been exclusively
confirmed in dipteran species: G.pallidipes, M.domestica, and possibly M.equestris. )t
has been proposed that GpSG(V and MdSG(V are phylogenetically related to baculoviruses, but have evolved in a very close association with their respective dipteran hosts. The hytrosaviruses share out of the baculovirus core genes
identified to date Jehle et al., , and are therefore, more distantly related to
baculoviruses than for instance the nudiviruses: Nudiviruses share of baculovirus
core genes Wang et al., . Nevertheless, these arguments appear to suggest a
Rationale
and
scope
of
this
thesis
The aim of this dissertation was to study the infectomics defined here as the functional and structural genomics and proteomics of GpSG(V. )t was conceptualized that the data obtained from these studies would be useful to develop novel, rationally designed strategies to manage GpSG(V infections in the laboratory colonies of
G.pallidipes. Chapter2 describes the dynamics and impacts of GpSG(V infection on
the productivity of G.pallidipescolony and modes of virus transmission. )n Chapter3,
GpSG(V strains circulating in wild populations of G.pallidipes are investigated.
Chapter4 investigates GpSG(V proteome, and correlates the viral ultra‐structure to
the protein composition, morphogenesis and cytopathology of the virus. )n Chapter5,
the role of tsetse saliva in the transmission of GpSG(V is investigated by determining
the secretome of asymptomatic and symptomatic G.pallidipes. Chapter6 investigates
the interplay between GpSG(V and the tsetse microbiome in trans‐generational virus
transmission. Chapter7 describes an essential advancement in the management of
GpSG(V in G.pallidipes colonies by modification of the invitro membrane‐feeding
regime traditionally used in tsetse mass – production facilities. Finally, Chapter8
Chapter
2
Dynamics
of
GpSGHV
transmission
in
laboratory
colonies
of
G.
pallidipes
Abstract
Tsetse flies Diptera; Glossinidae are naturally infected by the Glossinapallidipes
salivary gland hypertrophy virus GpSG(V . GpSG(V infection can either be asymptomatic or symptomatic, with the former being the most rampant in these colonies. Under yet undefined conditions, the asymptomatic state is triggered to a symptomatic state, leading to detectable salivary gland hypertrophy syndrome SG( that causes reproductive dysfunction and sometimes colony collapse. To gain a better
understanding of the impact of GpSG(V in G.pallidipes colonies, and to follow
development of SG( in the F progeny of symptomatic flies, progenies of tsetse flies reared under different conditions was examined. The results demonstrated that, whereas the F progeny of asymptomatic parents do not develop SG(, the F progeny of symptomatic females mated with asymptomatic males had a high SG( prevalence
rates % in male and % in females , and that these flies are sterile. Stress in the
form of high fly densities in holding cages flies/cage , and high temperatures
°C in the insectary lead to high mortalities and low productivity number of pupae/female . The number of viral particles secreted via saliva into blood during membrane feeding correlated with the infection statuses of the flies. After a single blood‐feeding event, asymptomatic and symptomatic flies release an average of
and viral genome copies/fly, respectively. Feeding the flies on fresh blood meals at
every feed for three fly generations significantly reduces the virus titres in these flies when compared with the viral titres in flies maintained under traditional feeding regime. The results of these studies allowed the initiation of colony management protocols aimed at minimizing the risk of horizontal GpSG(V transmission and enable establishment of SG( ‐ free colonies.
Introduction
)n many parts of sub‐Saharan Africa, trypanosomoses and the presence of tsetse are considered as major obstacles to the development of sustainable livestock production
systems and important root causes of hunger and poverty Dyck et al., ;
Feldmann etal., ; Jordan, . )t is generally accepted that control of the tsetse
vector is the most efficient and sustainable management for trypanosomoses (olmes
and Torr, ; Leak, ; Schofield and Kabayo, . The use of the sterile insect
technique S)T as a component of an Area‐Wide )ntegrated Pest Management AW ‐
)PM approach Klassen and Curtis, is a powerful fly control method as amply
demonstrated by eradication of Glossinaaustenifrom the )sland of Unguja, Zanzibar
Vreysen et al., . Efficient implementation of S)T depends on successful
maintenance of laboratory tsetse flies colonies to produce high quality males capable
of competing with wild males for mating with wild tsetse females (endrichs etal.,
.
)n laboratory colonies of G.pallidipes, infection by the salivary gland hypertrophy
virus GpSG(V can be either asymptomatic or symptomatic. Symptomatic infection is characterized by the salivary gland hypertrophy syndrome SG( , which can lead to
reproductive dysfunction and sometimes colony collapse Abd‐Alla et al., a .
)ncidence of asymptomatic infections can be high in both field and colonized tsetse
Odindo, . Asymptomatic infections are likely maintained through vertical
transmission, either via milk gland secretions or through gonadal tissues. The low virus titre in these asymptomatic flies does not cause measureable impacts on host fitness. Symptomatic infection is associated with testicular degeneration and ovarian
abnormalities Jura et al., ; Kokwaro et al., ; Sang et al., ; and
affects the development, survival, fertility and fecundity of naturally‐ or
experimentally‐infected flies Jura et al., ; Sang et al., . The incidence of
symptomatic infections is low zero – % in populations that often harbour high levels of asymptomatic infections. This chapter presents results of investigations into
the dynamics GpSG(V transmission in the laboratory colonies of G.pallidipes.Further,
data is presented on impact of stress high temperature and high population density on the prevalence of SG( syndrome in the colony productivity. The release of GpSG(V
particles via saliva into the blood during invitro feeding is quantified and correlated
Materials
and
methods
Tsetse
rearing
and
handling
Two G.pallidipes colonies were used in the study. A colony originating from pupae
collected in Tororo, Uganda in , colonized initially at the University of Leiden, The
Netherlands, and subsequently transferred to the )nsect Pest Control Laboratory
)PCL , Seibersdorf, Austria in Tororo colony Feldmann, a; Gooding etal.,
. A second colony was established at the Tsetse Fly Rearing and )rradiation Centre, Kality, Addis Ababa, Ethiopia from pupae collected near Arba Minch in the
period ‐ Arba Minch colony . Unless otherwise stated, experimental flies
were fed on heated, defibrinated bovine blood SVAMAN spol s.r.o., Myjava, ,
SLOVAK)A for ‐ min, three times/week using a membrane‐feeding technique
Langley and Maly, .
Two feeding protocols were used. A standard membrane feeding protocol, which is
routinely used in tsetse mass–production facilities Feldmann, a : )n this feeding
method, up to ten successive cages of flies were offered a blood meal on the same
membrane. A "clean blood feeding protocol" hereafter denoted as "clean
feeding" , in which each cage of flies was provided with a fresh blood meal at each feeding event. The clean feeding protocol was used to prevent flies from picking up viral particles from blood already used for feeding previous cages. Pupae produced
from sequential larviposition events were collected and incubated at °C until
emergence.
Diagnosis
of
GpSGHV
in
live
tsetse
To detect GpSG(V‐infected flies without dissection, a non‐destructive polymerase
chain reaction PCR method was used as preciously described Abd‐Alla et al.,
a . Briefly, total DNA was extracted from one intermediate leg excised from teneral newly ‐ eclosed, unfed flies collected within h post emergence, using ZR DNA genomic kit Zymo Research, California, USA according to supplier s instructions. The DNA was eluted in ‐μl elution buffer and stored at ‐ °C until further analyses. For PCR amplifications, . μl of the purified DNA was used as
template. The PCR reactions were performed to amplify a – bp fragment of the
coding sequence of GpSG(V ORF odv–e66 gene; GenBank accession No. EF ;
Abd‐Alla et al., . The following primers were used: GpSG(Vfwd – GCT TCA
GCA TAT TAT TCC GAA CAT AC ‐ , and GpSG(Vrev – GAT CCT GCT CGC GTA AAC
CA ‐ Abd‐Alla etal., a . The PCR amplification products were analysed on a
Quantification
of
GpSGHV
titres
in
individual
flies
Virus titres in individual legs or whole flies were assayed by qPCR as previously
described Abd‐Alla etal., b . Briefly, a calibration curve was set up for the qPCR
assay as follows: DNA extracted from purified virus was used to amplify the – bp
fragment of GpSG(V ORF described above, followed by purification of the PCR product using Q)Aquick PCR purification kit Qiagen . To ensure specificity and maximize qPCR efficiency, a pair of short specific primers for the GpSG(V ORF
flanked on the outside by the – bp fragment primers , was used to amplify a –
bp fragment of the gene using μl of the purified PCR product. The primers were as follows: ‐ QPCRFwd – CAA ATG ATC CGT CGT GGT AGA A ‐ , and QPCRRev – AAG CCG ATT ATG TCA TGG AAG G ‐ . The qPCR primers were designed to be as
short as possible nucleotides each to maximize PCR efficiency. The – bp PCR
product was purified, quantified by Nanodrop spectrometry, and the equivalent DNA
copies calculated according to standard protocols Sambrook etal., . To produce
the standard curves for estimation of viral titres in experimental samples by qPCR, ‐ fold serial dilutions of the purified DNA were made, and each standard was run in triplicate on the same ‐well qPCR plates with the test samples. Non – template controls water were included in the assay.
Effect
of
stress
on
SGH
prevalence
Male and female teneral flies were randomly selected from the Tororo colony, and
maintained in standard colony holding cages cm diameter x cm height at
different fly densities to flies per cage and mating ratios : and : ,
male: female Table1.
Table1: SetupoftheassaytodetermineeffectsofstressonSGHprevalence: Seven treatments, each replicated at least three times were set up at different fly densities – flies per cage , and at different male ♂: female ♀ ratios to assess the effects of stress on SG( prevalence in G.pallidipes colonies.
)t should be noted that in regular tsetse colonies, the normal fly density for the standard fly cages is flies at a male: female ratio of : . All the treatments were replicated at least three times. The replicate cages for each treatment were maintained at temperatures of °C or °C for days equivalent to blood meals . Fly mortality and productivity number of pupae per female were recorded. After days, all experimental flies were dissected to estimate occurrence of SG( symptoms.
Relationship
between
fly
productivity
and
SGH
prevalence
To investigate the relationship between SG( and fly productivity, teneral virgin
males and females were randomly selected from the Tororo colony, and separately maintained on clean blood meals until they reached sexual maturity. For laboratory –
bred G.pallidipesflies, maximum female receptivity occurs at – days of age, and
willingness of males to mate is maximal at – days of age Leegwater‐van der
Linden, . )t was not expected that the difference in sexual maturity between
males and females could negatively affect the experimental results: the first egg is usually retained and remains viable for several days after female maturation, and
would therefore, permit ovulation in response to the mating stimulus Wall, .
To compose parental G generations, single matings of ‐day ‐ old females and day ‐ old males were performed in individual plastic tubes cm diameter and cm height with netting on top and bottom for feeding and pupae collection . Each tube containing the experimental flies was numbered to identify the individual mating partners. Flies were allowed to mate for h, after which the males were removed and dissected to assess occurrence of SG(. Females were offered clean blood meals
until they produced three F larviposition pupae, or until days post mating,
whichever was the earlier. Subsequently, the female flies were also dissected to assess SG(. The F pupae from individual females were collected in plastic tubes and labelled as described above for the parents. Based on the parental SG( status, the F pupae were divided into four groups, those from ) asymptomatic male and female parents, )) asymptomatic male and symptomatic female parents, ))) symptomatic male and asymptomatic female parents, and )V symptomatic male and female parents. Pupae
from these four groups were incubated at °C for days or until emergence.
)ndividual pair matings were made between F males and females within each group of pupae and the males and females were treated as for the G . The F pupae were collected from each individual female fly and incubated at °C until emergence.
Detection
and
quantification
of
GpSGHV
in
blood
meals
after
feeding
Symptomatic and asymptomatic tsetse flies eight flies in each category; replicated
three times were fed individually on ~ µl clean blood for ‐ min. Only flies
that were fully engorged fully fed at the end of the feeding event were further analysed. After feeding, the blood that remained under the feeding membranes was
collected for subsequent DNA extraction. For negative control, ~ µl of the clean
blood was sampled prior to each feeding event. Total DNA was extracted from the collected blood using the DNeasy kit Qiagen following the supplier s instructions, and virus was detected by the end‐point PCR protocol as described above. Viral titres in the samples were quantified by qPCR.
Controlled
feeding
of
tsetse
on
GpSGHV
‐
contaminated
blood
and
analysis
of
virus
particles
secreted
via
saliva
Teneral G.pallidipes flies were screened by end‐point PCR to determine GpSG(V
infection status. Sixteen symptomatic flies were selected, maintained individually in
numbered plastic tubes, and fed on ~ μl clean blood meals. After feeding, the
blood remaining under the membranes was collected, thoroughly mixed and divided
into two aliquots ~ μl each ; one aliquot was used for DNA extraction and
subsequent qPCR analysis, and the other was used to feed asymptomatic PCR ‐ negative flies. The PCR ‐ negative flies were divided into groups each composed of
flies , and given one, three, five or seven successive GpSG(V‐contaminated blood meals. After receiving the respective number of blood meals, the first three fly groups were offered clean blood meals to bring the total to seven blood meals. Negative control flies received seven meals on clean blood. All fly groups were offered an additional eighth clean blood meal, and the blood residue after this final feed was analysed by end‐point PCR to detect GpSG(V.
Impact
of
clean
feeding
on
GpSGHV
titres
Parental G generation for the bioassay consisted of male and female teneral flies was randomly selected from the Tororo colony. The flies were maintained on clean feeding for sixty days, and the pupae were collected. To determine GpSG(V titres at the start of the assay, twenty‐four G flies six males and eighteen females were randomly sampled and stored at – °C for qPCR analysis. After incubation at °C, the flies emerging from the F pupae were subsequently mated and maintained
as above, and F pupae were collected. The F and F adults were then sampled and
Statistical
analysis
To compare means of GpSG(V titres and SG( prevalence rates between the experimental fly groups, statistical analysis was performed according to Sokal and
Rohlf Sokal and Rohlf, . To find out actual significant differences between the
treatments groups , analysis of variance ANOVA was followed by Tukey s (SD honestly significant difference Test for unplanned "a posteriori" comparisons of
means and Student s t ‐ test for regression coefficients. Pairs of proportions were
compared using the likelihood ‐ ratio G test.
Results
This study was designed to investigate; i the influence of fly density and insectary conditions on GpSG(V titres and SG( prevalence rates, ii the interplay between SG( prevalence, fly mortality and productivity, and iii the dynamics of acquisition of GpSG(V particles released by infected flies via saliva during membrane.
Diagnosis
of
GpSGHV
in
live
tsetse
GpSG(V infection statuses in the fly colony could be divided into three categories:
negative, slightly positive and strongly positive Figure1 .
Dissection of flies from each category showed that, whereas the PCR ‐ negative and slightly positive flies did not reveal any detectable SG( symptoms, % of the flies in the strongly positive group had overt SG( symptoms.
Prevalence
of
GpSGHV
in
Seibersdorf
and
Kality
fly
colonies
Between and , regular dissection of batches of ~ flies from the Tororo
colony has shown a stable SG( prevalence less than % . (owever, SG( prevalence
Figure 1: PCR diagnosis of GpSGHV infections in
G.pallidipes: DNA was
extracted from one
mesothoracic leg from
in the Arba Minch colony maintained at the Tsetse Fly Rearing and )rradiation Centre,
Kality, Addis Ababa, Ethiopia, showed significantly higher SG( rates of . % in ,
. %, in and % in (Figure2) as compared to those obtained in the
Tororo colony , P< . ; , P<< . ; , P< . .
The data for and were obtained from flies dissected at Kality and the data
for were obtained from flies dissected at )PCL, Seibersdorf, which had emerged
from pupae originating from Kality. The high SG( prevalence has been accompanied by a decline in the size of the Kality colony.
Figure2:PrevalenceofSGHsyndromeinG.pallidipescolonies: The figure shows differences in the SG( prevalence in the Tororo G.pallidipescolony maintained at the )PCL, Seibersdorf, Austria, and the Arba Minch colony maintained at Kality, Addis Ababa, Ethiopia likelihood ratio test, * P< . , ** P< . , ***
P< .
Effects
of
temperature
and
fly
density
on
prevalence
of
SGH
syndrome
A high proportion of G.pallidipes flies were asymptomatic. Further, whereas at °C
the number of flies per cage and the sex ratio had a limited effect on mortality, a
significant increase in mortality was observed in flies reared at °C Figure3 ,
)n the flies that were maintained at °C, female productivity progressively decreased with an increase in fly density regardless of the sex ratio. The productivity of females was significantly reduced at °C, with the lowest productivity at the highest density
of flies per cage Figure4.
Although SG( prevalence varied from ‐ . % depending on treatment, there was no clear correlation between fly density or rearing temperature and the percentage of flies exhibiting detectable SG(. (owever, it cannot be excluded that the dead flies included a higher proportion of flies with SG( and that the symptomatic flies showed
lower fecundity than the asymptomatic flies Jaenson, .
Figure3:Impactsofstressonflymortalities:The figure shows effects of fly density and temperature on mortality on flies sampled from the Tororo G.pallidipescolony : , cages with sex ratio male: female flies; : , cages with sex ratio male: female fly .
Relationship
between
SGH
and
tsetse
productivity
Correlation of fly productivity with SG( syndrome is presented in Table2. Dissection
of flies males and females at the end of the G showed an average SG(
prevalence of . % in both sexes. From the matings, were classified in
