IAEA-314-D4-RC-1048.1 LIMITED DISTRIBUTION

WORKING MATERIAL

IMPROVING SIT FOR TSETSE FLIES THROUGH RESEARCH ON THEIR SYMBIONTS AND PATHOGENS

FOURTH RESEARCH COORDINATION MEETING

ORGANIZED BY THE JOINT FAO/IAEA DIVISION OF NUCLEAR TECHNIQUES IN FOOD AND AGRICULTURE

25 th – 30 th March, 2012 Vienna, Austria

NOTE The material in this document has been supplied by the authors and has not been edited by the IAEA. The views expressed remain the responsibility of the named authors and do not necessarily reflect those of the government of the designating Member State(s). In particular, neither the IAEAA nor any other organization or body sponsoring the meeting can be held responsible for any material reproduced in this document.

Contents 1.INTRODUCTION ...... 4 1.1.T SETSE AND THE DISEASE ...... 4 1.2 TRYPANOSOMOSIS CONTROL AND THE SIT ...... 4 1.3. CONSTRAINTS AND TARGETS FOR IMPROVED EFFICIENCY OF TSETSE SIT ...... 5 1.4 TSETSE SYMBIONTS ...... 5 1.5 TSETSE PATHOGENS ...... 6 2. CURRENT STATUS ...... 7 2.1. TSETSE SYMBIONTS ...... 7 2.2. TSETSE FLY PATHOGENS ...... 11 2.2. TSETSE FLY PATHOGENS ...... 11 3. INDIVIDUAL ACHIEVMENTS DURING THE CRP ...... 14 4. LOGICAL FRAMEWORK ...... 45 5. AGENDA ...... 51 6- LIST OF PARICIPANTS ...... 55 ANNEX I: WORKING PAPERS ...... 59 Page 3

Executive Summary

Major efforts have been expended by a number of international scientists in a focused and coordinated manner to (i) elucidate the cause of the collapse of tsetse fly colonies and reduced fly yields in tsetse fly production facilities for SIT, and (ii) to put into practice an action plan to solve the problem by understanding the biological basis of the symptoms in the flies associated with the collapse. Through a coordinated action, it has been found that a virus is the culprit causing reduced fecundity and that the disease is characterized by the hypertrophy of the salivary glands, hence the name, salivary gland hypertrophy virus (SGHV). The virus genome has been sequenced and compared to a related virus in the house-fly. SGHV is vertically and horizontally transmitted. The incidence of the disease in the field throughout Sub-Saharan Africa ranges from 0-15%, but approaches 100% in laboratory reared colonies. Through this CRP, tsetse’s physiology including fecundity has been shown to depend upon fitness of its symbiotic fauna. Correlations and interactions between the presence of virus, disease symptoms, and the occurrence of bacterial symbionts ( Wigglesworthia , Sodalis and Wolbachia) has been explored. Strategies have been designed and partially validated to mitigate / control / eliminate the disease. These strategies are based on: (i) monitoring viral loads for colony quality control; (ii) blocking transmission using specific antibodies, and/or clean feeding practices; (iii) blocking virus replication by applying specific inhibitors of virus replication. Strategies have also been designed to: (i) monitor prevalence and loads of tsetse symbionts and pathogens; (ii) augment current feeding regiments to improve tsetse’s fecundity; (iii) improve the application of SIT by harnessing tsetse’s symbionts to develop pathogen resistant fly lines and by introducing natural sterility. The outcome of the research efforts provided the fundamental knowledge required on tsetse’s microbiome and SGHV to develop a solid and robust strategy to prevent colony collapses caused by SGHV. This will have a major impact on large-scale production facilities for SIT. Further experimentation is required, including efficacy testing, to make this strategy robust, reliable and applicable for large-scale production facilities. The anticipated outcome will help to improve SIT applications, which will be more efficacious and cost-effective. The program in addition has resulted in extensive capacity building and technology transfer to disease endemic countries (DECs). Concerted action has enabled the involvement of all participants in the research to ascertain the application of a well-orchestrated strategy and production process to secure high fly yields in the future for more sustainable SIT applications.

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1.Introduction 1.1.Tsetse and the Disease Tsetse flies (Diptera: Glossinidae) are the cyclical vectors of the trypanosomes, which cause human African trypanosomosis (HAT) or sleeping sickness in humans and African animal trypanosomosis (AAT) or nagana in animals. It is conservatively estimated by the World Health Organization (WHO) that there are currently 10,000 – 45,000 cases of human African trypanosomosis (HAT) with 60 million people at risk in 36 countries covering ~40% of Africa (almost 10 million km²). After a devastating epidemic in the early 20 th century when a million people died of HAT the disease almost disappeared by the 1960’s from Africa. HAT has a disease burden of 1.47 million disability adjusted life years (DALY). Given that the disease affects hard to reach rural populations and that there is a lack of active surveillance in war-torn areas, the disease prevalence numbers are undoubtedly a gross underestimation. In addition to direct losses of human life and the financial cost of control, African trypanosomosis affects:

• Human health, through protein deficiencies caused by shortage of meat and milk • Livestock production, since it causes morbidity and mortality • Agricultural production, through the lack of draught animals and manure • Rural economy, by preventing integrated agriculture and livestock production • National economy, through essential import of living animals and their products • Environment, though the use of insecticides.

The tsetse and trypanosomosis (T&T) problem is a key obstacle for rural development. About 50 million cattle and tens of millions of small ruminants are at risk from AAT. T&T are the major factors preventing the establishment of sustainable agricultural systems in sub-Saharan Africa. The Programme Against African Trypanosomiasis (PAAT) estimates that AAT causes approximately 3 million cattle deaths per year. Farmers are required to administer approximately 35 million doses of costly trypanocidal drugs. Unfortunately, many of these chemicals fail because of development of resistance by the parasites. Direct losses in meat production and milk yield and the costs of programmes that attempt to control trypanosomosis are estimated to amount to between US$600 million and $1.2 billion each year. In the absence of the AAT problem, a family that is currently dependent on manual labour alone could use draught animals and thus increase its income from agricultural work by 45 percent per unit of land and by 143 percent per unit of labour. If one includes this potential in livestock and crop production, trypanosomosis is estimated to cost sub- Saharan Africa US$4 billion or more each year, equivalent to one-quarter of the area's total livestock production. 1.2 Trypanosomosis Control and the SIT Human sleeping sickness is a zoonosis caused by the protozoan Trypanosoma brucei rhodesiense in East Africa and T. b. gambiense in West and Central Africa. The nagana causing related trypanosomatids T. vivax and T. congolense are major pathogens of cattle and other ruminants, while T. simiae causes high mortality in domestic pigs and T. brucei affects all livestock. Vaccines are not available and are unlikely to be developed due to the antigenic variation in the trypanosome in the mammalian host. The drug treatment of HAT is in a perilous state and relies on old, often dangerous drugs and resistance is becoming an increasing problem. In contrast, disease control, via control of the tsetse vector, has been found to be highly effective. Current vector control efforts, which depend on trapping or killing the tsetse flies with insecticides, have been difficult to sustain at the local community level.

The successful eradication of the tsetse fly Glossina austeni on Unguja island of Zanzibar by means of an area-wide integrated pest management programme concluding with the release of sterile flies Page 5 provided new opportunities and stimulated interest to expand the technology to large areas on mainland Africa. Encouraged by this success, in July 2000, at a summit in Lomé, Togo, the African Heads of State and Government passed a resolution (AHG/Dec.156 XXXVI), giving birth to the Pan African Tsetse and Trypanosomosis Eradication Campaign (PATTEC).

Several UN organizations, including WHO, the Food and Agricultural Organization (FAO) and the International Atomic Energy Agency (IAEA) subsequently passed resolutions supporting the PATTEC initiative. In May 2002 PATTEC and PAAT jointly developed criteria for the identification of priority areas for T&T intervention and for related sustainable agriculture and rural development. Some priority areas for T&T intervention and rural agriculture and livestock development in eastern, western and southern Africa were screened according to these criteria, and specific national / sub-regional strategies were developed and necessary national and international funding could be generated, including grants and soft loans from the African Development Bank. 1.3. Constraints and Targets for Improved Efficiency of Tsetse SIT Despite the successful use of sterile tsetse flies in area-wide integrated vector control programmes, several constraints needed to be addressed so as to enhance efficiency and expand areas where sterile males can be used. Some of these constraints are a function of the interaction of the tsetse fly with its symbionts and pathogens and were the targets for this CRP. 1.3.1. Colony performance of Glossina pallidipes . This very important species of tsetse and the target for several area-wide programmes in eastern Africa experiences unpredictable problems in mass rearing performance which seriously compromises effective implementation of field releases. This reproductive anomaly is due to the presence of a virus that causes salivary gland hypertrophy in a proportion of the infected individuals. 1.3.2. Trypanosome resistant strains for release. As the sterile tsetse flies for release are potential vectors, specific measures have to be taken to ensure that they do not contribute to disease transmission following release. In practice this means removing the long-lived females from the release and providing two blood meals, containing a trypanocide, to the males before release. Manipulation of the symbionts present in tsetse can provide a direct means to achieve disease resistant (refractory) strains for release. 1.3.3. Natural incompatibility . Natural incompatibility between populations of the same species can provide an additional tool for vector control programmes. One class of symbionts present in tsetse has been shown to be responsible for this incompatibility in many insect species. However, their possible role in incompatibility between tsetse populations remains to be established. This type of incompatibility can also be used to spread useful genes into populations. 1.3.4. Suppression . Suppression of target tsetse populations before release of sterile insects is essential and methods range from aerial spraying of insecticides to community based use of traps and targets, which also can incorporate insecticides. Fungal pathogens spread by tsetse themselves (autodissemination) may offer an alternative to the use of insecticides.

1.4 Tsetse Symbionts Tsetse flies are unusually dependent on their microbial flora for providing nutrients that are not present in their restricted diet of vertebrate blood. To date three organisms that are maternally transmitted have been identified in tsetse field populations and laboratory colonies and the genomes of these three organisms have been sequenced. 1.4.1. Wigglesworthia . This is an intracellular obligate microbe harboured by all tsetse species. Its elimination from tsetse results in significantly reduced host fecundity as well as deficiencies in immune maturation. Its genome sequence indicates extended similarities to other primary obligate symbionts of insects. Its symbiotic role in tsetse appears to be nutritional in nature, as evidenced by many vitamin biosynthetic pathways its genome encodes. Knowledge on

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Wigglesworthia products and transmission biology to progeny is central for successful maintenance of tsetse colonies. 1.4.2 . Sodalis. This organism resides in the midgut where it is in close proximity to the site of trypanosome development. It has been possible to establish in vitro culture conditions for Sodalis , introduce and express foreign genes and to reintroduce the modified symbionts into tsetse. Ability to express antitrypanosomal gene products in Sodalis can result in tsetse strains that are unable to transmit trypanosomes and will enhance the efficacy of SIT applications and in addition can result in novel methods through a population replacement strategy. 1.4.3. Wolbachia . These organisms are obligatory intracellular and maternally transmitted bacteria that infect many insect and nematode species. These bacteria cause a number of reproductive alterations such as parthenogenetic development, overriding of chromosomal sex determination to convert genetic males into functional females, killing male embryos at early developmental stages, and cytoplasmic incompatibility (CI). Many tsetse populations have been shown to be infected with Wolbachia strains and indirect evidence for cytoplasmic incompatibility exist. Direct demonstration of the ability of tsetse Wolbachia strains to induce cytoplasmic incompatibility is necessary. In addition, Wolbachia has been shown to protect insect hosts against viral infections as well as limit infections with pathogens such as Dengue, Chikungunya and Plasmodium. The influence of tsetse Wolbachia on the prevalence of pathogens such as GpSGHV and trypanosomes needs to be investigated.

1.5 Tsetse Pathogens Pathogens of tsetse interact with SIT in two ways, one negative and one positive. Firstly the presence of a virus can compromise the productivity of laboratory colonies and so interfere with effective mass rearing, and secondly fungal pathogens may be used as a means to suppress tsetse populations prior to the release of sterile males. 1.5.1 Salivary gland hypertrophy virus (SGHV). The appearance of abnormal mortalities due to virus infection is well documented in mass rearing of medically and/or economically important arthropods. In Glossina pallidipes SGHV is responsible for a typical teratogenic symptom of salivary gland hypertrophy (SGH) affecting both males and females and leading to infertility in the affected insects. The virus has been implicated in the collapse of a colony established from Ethiopia and it is also associated with unpredictable reductions in colony productivity. The virus replicates in the salivary gland, the female milk gland, midgut ovaries and testes and is known to be transmitted vertically (transovum transmission) and presumably also sexually (veneral transmission). Virus specific primers targeting the GpSGHV are available for diagnosis and generated primers will be prepared. 1.5.2. Fungus. Although entomopathogenic fungi can cause lethal or debilitating diseases in insects, little information is available concerning entomopathogens associated with tsetse flies in the field. Studies have shown that many species of tsetse flies are susceptible to fungal infection in the laboratory. However, the major obstacle to utilization of a fungal pathogen has been the difficulty of its application in the field. Recently a contamination device (CD) was developed where attracted flies are contaminated with spores before they return to the environment where they may contaminate other flies during mating. This results in auto-dissemination of the fungus. 1.5.3 Glossina iflavirus. A potential picorna-like virus was detected in a cDNA library of G. morsitans morsitans . From the sequence information it appears that the virus is different from the Deformed Wing Virus (DWV) group, but a putative member of the Iflavirus family (Iflaviridae). So far the virus, provisionally named Glossina iflavirus, has not been physically detected (as 30 nm particles) in salivary gland tissue and no particular pathology has been noticed. The Glossina iflavirus could be a potential control agent of tsetse flies. DWV is the major cause of colony collapse in bees. Page 7

2. Current Status 2.1. Tsetse symbionts. Tsetse flies are unusually dependent on their microbial flora for providing nutrients that are not present in their restricted diet. To date three microorganisms that are maternally transmitted have been identified in tsetse populations and colonies, namely Wigglesworthia, Sodalis and Wolbachia . To analyze the precise tissue distribution of the three endosymbiotic bacteria and to infer the way by which each symbiotic partner is transmitted from parent to progeny, Aksoy and Heddi have used Fluorescence in situ Hybridization (FISH) methodology that allows the specific screening of bacteria through the design of specific 16S rDNA probes. Results showed that bacteriocytes are mono-infected with Wigglesworthia , while both Wigglesworthia and Sodalis are present in the milk gland lumen. Sodalis was further seen in the uterus, spermatheca, fat body, milk and intracellular in the milk gland cells. Contrary to Wigglesworthia and Sodalis, Wolbachia were the only bacteria infecting oocytes, trophocytes, and embryos at early embryonic stages. Furthermore, Wolbachia were not seen in the milk gland and in the fat body. This work further highlights the diversity of symbiont interactions in multipartner associations and supports two maternal routes of symbiont inheritance in the tsetse fly: Wolbachia through oocytes, and, Wigglesworthia and Sodalis by means of milk gland bacterial infection at early post-embryonic stages (Balmand et al., 2012) . Wigglesworthia : The enteric Wigglesworthia glossinidia is an obligate mutualist microbe harboured by all tsetses. Wigglesworthia resides intracellularly within differentiated epithelial cells (bacteriocytes), in a special bacteriome in the anterior part of the midgut. The association between Wigglesworthia and its tsetse host is apparently ancient (50-80 millions years old) and phylogenetic studies show that Wigglesworthia species display concordance with their insect host species, a phenomenon indicative of strict vertical transmission of the symbiont from generation to generation. The genome sequence of two Wigglesworthia species is available from the host species G. m. morsitans and G. brevipalpis , respectively. Both genomes indicate extended similarities with other primary obligate symbionts of insects, which similar to tsetse live on strictly restricted diets. Wigglesworthia genomes show a drastic reduction in size to about 700 kb and exhibit a high A+T bias of about 80%. The elimination of Wigglesworthia via antibiotic supplementation of tsetse’s diet results in loss of host fecundity. Its symbiotic role in tsetse appears to be nutritional in nature, as evidenced by many vitamin biosynthetic pathways its small genome encodes. In addition, when larvae develop in the absence of Wigglesworthia , emerging adult progeny are deficient in immune responses, in particular in cellular immune pathways. The small genome of Wigglesworthia has retained the ability to synthesize a functional flagellum. Gene expression studies indicate that the flagellum is expressed in the milk gland tissue suggesting a role for flagella functions in the transmission process. The Wigglesworthia morsitans genome has revealed several functional capabilities that are lacking in Wigglesworthia brevipalpis . One of these differences suggests that W. morsitans has the ability to encode charismate leading to the production of folate. The absence of genes in these pathways suggests that folate biosynthesis may not possible by W. brevipalpis . Given that protozoan parasite African trypanosomes are auxitrophs for folate, the synthesis of folate by W. morsitans may contribute to the well-documented enhanced vector competence phenotype of its host G. morsitans species. Wigglesworthia is not detected in the germ-line tissues of tsetse but is acquired by tsetse’s intrauterine progeny through the maternal milk gland secretions.

Sodalis : The second organism, facultative Sodalis, is harboured in all tsetse flies in laboratory lines and lives in close proximity in the midgut to where trypanosomes develop. Interestingly, closely related symbionts have been identified in insects from different taxa such as hippoboscid flies (Diptera), weevils (Coleoptera) and chewing lice (Phthiraptera). The genome sequence of Sodalis is also available, further mediating functional studies into Sodalis’ biology in the tsetse host. Similar to Wigglesworthia , Sodalis is acquired by the intrauterine larva through the mother’s milk gland secretions as has been documented in this CRP. Studies where Wigglesworthia

Page 8 symbiont has been selectively eliminated have led to loss of Sodalis in subsequent generations, indicating the presence of cooperative interactions between the obligate symbiont and Sodalis An interesting hypothesis has focused on the functional role of Sodalis for trypanosome transmission. The presence of Sodalis has been implicated in enhancing tsetse’s vectorial capacity. Individual flies that harbour greater Sodalis densities have been suggested to be more susceptible to trypanosomes. In addition to density effects, it is also possible that different Sodalis genotypes may confer greater transmission ability to the host. These inferences can be evaluated further in populations both for density effects and for potentially different genotypes that may exist in natural populations to confer enhanced parasite transmission ability to host insects. In a previous work, a large tsetse fly sampling campaign had been conducted in Bipindi and Campo that are two historical sleeping sickness foci located in the south of Cameroon. More than four thousand flies had been trapped. For each fly, we have determined the species to which it belonged, the presence/absence of S. glossinidius , and the presence/absence of trypanosome infection. In case of infection, the species and subspecies of the infecting parasite have been identified, and the prevalence of each of them was calculated. The results evidenced large differences between the population of flies from Campo and that from Bipindi (Farikou et al., 2010). So, the next step was to identify possible genetic diversity in symbiont populations. We first try to apply microsatellite genotyping method on insectary flies using four markers (Farikou et al., 2011a). After we have verified that the method was successful, we have applied the same method on field tsetse flies from Cameroonian foci allowing detection of Sodalis diversity in field flies (Farikou et al., 2011b). The Sodalis endosymbiont has been proposed as a potential delivery vehicle of proteins/peptides to control trypanosome parasite development in the tsetse fly, an approach known as paratransgenesis. One application of this approach for SIT would be the engineering of released tsetse fly lines that are unable to transmit trypanosomes. For this, gene products that interfere with trypanosome development have to be expressed and released by Sodalis to the outer environment without reducing the fitness of the tsetse host or its symbiont. Expression of heterologous proteins in cultured Sodalis has been achieved. A functional twin-arginine (TAT) translocation pathway in Sodalis to export active heterologous proteins to the periplasm has been demonstrated (De Vooght et al., 2011). In addition, it was shown that also the pelB leader peptide was successful in directing the export of functional anti-trypanosome single domain antibody (Nanobody ®) to the periplasm of S. glossinidius resulting in significant levels of extracellular nanobody release. These S odalis strains are not affected in their growth compared to the wildtype Sodalis , suggesting that they may be competitive with endogenous microbiota in the midgut environment of the tsetse fly. These data are the first demonstration of the expression and extracellular release of functional trypanosome- interfering proteins in S. glossinidius , further supporting its use as a paratransgenic platform organism (De Vooght et al., 2012).

Wolbachia : The third symbiont Wolbachia is an obligatory intracellular and maternally transmitted bacterium that infects many arthropod and filarial nematode species. These bacteria cause a number of reproductive alterations such as parthenogenetic development, overriding of chromosomal sex determination to convert genetic males into functional females, killing male embryos at early developmental stages, and cytoplasmic incompatibility (CI). CI, which has been documented in all major insect orders, can be either uni- or bidirectional. Unidirectional CI is typically expressed as embryonic lethality when an infected male is crossed with an uninfected female. Bidirectional CI occurs in crosses between infected individuals harbouring different strains of Wolbachia , that is, strains with different CI properties. In most insects, the expression of CI is lethal to the developing embryo. The molecular mechanism, which is responsible for the induction of CI, has not yet been unravelled. However, Wolbachia and Wolbachia -induced CI have been proposed as a tool to control pests and diseases.

In the frame of this context and CRP, we aimed at: (a) the detection and characterization of Wolbachia infections in both laboratory and natural populations of tsetse flies, (b) the possible Page 9 correlation between Wolbachia and salivary gland hypertrophy virus infections and (c) Wolbachia - induced CI in infected Glossina lines and its potential to be used for the control of tsetse flies and trypanosomosis.

A specific 16S rRNA PCR assay (Doudoumis et al., 2012) was used to detect the presence of Wolbachia in a total more than 5000 specimens of eleven different Glossina species (G. m. morsitans, G. m. centralis, G. m. submorsitans, G. austeni, G. brevipalpis, G. pallidipes, G. p. palpalis, G. p. gambiensis, G. fuscipes fuscipes, G. tachinoides and G. medicorum from laboratory stocks and natural populations originating from 15 African countries. Wolbachia was detected in all Glossina species while the prevalence varied. The tsetse Wolbachia strains were characterized based on the Multi Locus Typing System (MLST) and the wsp gene (outer surface protein) including an HRM approach. In total, we detected multiple allelic profiles or Sequence Types (ST), wsp alleles and WSP HVR profiles. The available sequence information was used for phylogenetic analysis, which suggested that Glossina species have experienced repeated invasions by Wolbachia strains.

In addition, multi-copy gene marker systems were developed and optimized that target Wolbachia transposons (IS elements) of tsetse flies in order to improve symbiont detection limits appropriate for large scale PCR screening. Furthermore, so-called wsp-Blot-PCR techniques were successfully adopted for tsetse, which furthermore enhance detection limits of this symbiont by three orders of magnitude. This combined high-resolution PCR approach uncovered the existence of so-called low- titer Wolbachia infections in natural and lab colonies of tsetse hosts that were earlier reported as uninfected by using standard PCR techniques. Finally, by targeting hyper-dynamic intergenic regions of Wolbachia that are composed of variable numbers of tandem repeats (VNTRs), even closely-related strains could be easily distinguished by fragment size polymorphism in multi- infected tsetse flies as well as symbiont copies could be designated that have been translocated on tsetse host chromosomes.

We also detected multiple horizontal transfer of Wolbachia genomes into the chromosomes of G. m. morsitans and, most likely, in other Glossina species too. The genome sequence of the cytoplasmic wGmm strain and of the chromosomal insertions was determined. The majority of horizontally transferred genes are pseudogenized. Ankyrin genes, prophages and insertion sequences were also determined in both cytoplasmic and nuclear Wolbachia genomes. Interestingly, a negative correlation between the presence of Wolbachia and the salivary gland hypertrophy virus was observed. A major finding of this CRP was that the presence of Wolbachia in G. m. morsitans is associated with the strong expression of cytoplasmic incompatibility, which reached the levels of 100% (Alam et al., 2011). This finding suggests that tsetse Wolbachia symbionts could be used for the control of the insect vector and of trypanosomosis. This can be done with three potential approaches: (a) to use Wolbachia -induced CI as a population suppression mechanism in a way analogous to SIT (Apostolaki et al., 2011; Laven, 1967; Zabalou et al., 2009; Zabalou et al., 2004); (b) to use Wolbachia -induced CI as a spreading/replacement mechanism for desired phenotypes. Infected females have a reproductive advantage over uninfected females, since they can mate with both uninfected and infected males, thus their genotype invades populations in nature (Bourtzis, 2008; Bourtzis and Robinson, 2006; Bourtzis and O’Neill, 1998; Brelsfoard and Dobson, 2009; Dobson et al., 2002; Rasgon, 2007, 2008; Sinkins and Gould, 2006) and (c) Wolbachia -induced CI can also be used to drive trypanosome resistant paratransgenic tsetse into natural populations to replace their parasite-susceptible counterparts (Alam et al., 2011). Based on the fact that all symbionts are maternally transmitted to the progeny, resistance genes expressed by Sodalis can be propagated by Wolbachia induced CI , if complete transmission of Sodalis and Wolbachia can be ensured (Aksoy et al., 2008; Aksoy and Weiss, 2007; Rio et al., 2004).

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Development of lines that are symbiont-cured . We evaluated the effect of continuous per os treatment of fertile females with the tetracycline and ampiciline based antibiotics and yeast extracts or vitamins mixture diets by measuring the total number of larvae deposited per group over the course of three deposition cycles. To determine if the tetracycline and ampicilin antibiotics have a prolonged effects on the process of metamorphosis within the larvae or pupae, the hatching rate of progeny was determined in these groups. At the same time we measured the influence of antibiotic and yeast treatment on fly lifespan. We measured the effect of tetracycline, tetracycline and yeast, ampiciline, ampicilin and yeast ; only yeast and different vitamine mixture treatment on pupae production. Under optimum conditions the first gonotrophic cycle takes about 20–22 days for development from egg to parturition. In subsequent gonotrophic cycles females produce a larva every 9 to 11 days. Ampicilin treatment does not reduce fecundity since it does not damage Wigglesworthia resident within bacteriocytes in the midgut, unlike tetracycline, which clears all bacteria including Wigglesworthia and Wolbachia and induces sterility. Accordingly, ampicillin- receiving flies remained fecund while tetracycline receiving flies were rendered sterile. Yeast extract (10% w/v) provisioning of the blood meal rescued fecundity of the females receiving tetracycline to similar levels as that of wild type and ampicillin receiving flies (65%, 55% and 64% over the first gonotrophic cycle and 53%, 58% and 49% over the second gonotrophic cycles, respectively). However, yeast provisioning at 10% w/v had a cost on fecundity when compared to flies maintained on normal blood meals, (92% versus 55% over the first gonotrophic cycle and 92% and 58% over the second gonotrophic cycle, respectively). Nevertheless, yeast supplementation was able to rescue the tetracycline-induced sterility to levels comparable to those observed for Gmm Wt receiving yeast or ampicillin supplemented blood meals, respectively. Thus yeast supplemented dietary regiment allowed us to develop two lines to analyze the functional role of Wolbachia symbionts in tsetse biology; one lacking all symbionts ( Gmm Apo ) and another lacking Wigglesworthia but still retaining Wolbachia and Sodalis (Gmm Wig−). The Gmm Apo progeny resulting from the first and second depositions of tetracycline treated mothers were tested for the presence of Sodalis, Wigglesworthia and Wolbachia by a bacterium-specific PCR-assay. The PCR- assay demonstrated the absence of all three symbionts as early as the first deposition in both the male and female Gmm Apo adults. The absence of Wolbachia from the reproductive tissues of Gmm Apo females was also verified by Fluorescent In Situ Hybridization (FISH) analysis. In contrast, Wolbachia was present in egg chambers during both early and late developmental stages in Gmm Wt females. Longevity of F1 Gmm Apo females was compared to that of Gmm Wt adults maintained on the same yeast-supplemented blood meal over 40 days (two-gonotrophic cycles). No difference (X 2 = 0.71, df = 1, P = 0.4) was observed in survivorship comparisons between the two groups. The second line ( Gmm Wig−) generated from ampicillin treated females still retained their Wolbachia and Sodalis symbionts, while lacking both Wigglesworthia populations as evidenced by FISH and PCR amplification analysis. When maintained on yeast-supplemented blood, this line (similar to Gmm Apo ) also did not display any longevity differences from the Gmm Wt adults sustained on the same diet. Through understanding the mechanisms by which tsetse endosymbionts potentiate trypanosome susceptibility in tsetse, it may be possible to engineer modified endosymbionts which, when introduced into tsetse, render these insects incapable of transmitting parasites. In our study we have assayed the effect of two different antibiotics on the endosymbiotic microflora of tsetse Glossina genus. We showed that the antibiotics, ampiciline and tetracycline, have a different impact on tsetse fecundity, pupal emergence, kortality, efectively rendering these insects sterile. Using the yeast extracts and different vitamine mixtures partially eliminate the antibiotic efect. Base on our results we can consider the yeast diet as the most appropriate to sustain optimal colony of endosymbiont-free flies.

Symbiotic influences on host reproduction : Data have been obtained on spatial/temporal variation in the reproductive behaviour of G.fuscipes fuscipes and G. pallidipes. In particular the occurrence of Page 11 female mating and remaiting has been assessed in two Uganda populations and Kenya, respectively. Different mating and remating estimates provide evidence that remating is a frequent behaviour in the wild and what is more, females store sperm from multiple males, which are potentially available for insemination. In these two populations Wolbachia infection frequency is high.

2.2. Tsetse Fly Pathogens 2.2. Tsetse Fly Pathogens The majority of research on the Glossina pallidipes salivary gland hypertrophy viruses (GpSGHVs) has been conducted under objective 2 to understand and manage the tsetse fly-SGHV interactions in lab populations. The first of the five verifiable indicators for this objective was the genome analysis of GpSGHV isolated from the Uganda G. pallidipes colony, maintained since 1979 at Seibersdorf. The collaborative research effort of the IAEA laboratories at Seibersdorf and at the University of Montpellier have produced and published a full-length sequence of GpSGHV (Abd-Alla et al. , 2009). In addition, the genome of the GpSGHV isolated from the Ethiopian strain of a G. pallidipes colony also has been sequenced. The purpose was to: 1) determine whether or not this sequence differed significantly from that of the Uganda strain, 2) compare the assembly and annotation of the two genomes, and 3) correlate sequence differences between the two GpSGHV isolates with the collapse of the Ethiopian colony. In fact, the two viruses share 98.7 % identity, their overall organization is almost identical, and 80 % of their ORFs have the same best blastp, strongly supporting the conclusion that they are two GpSGHV strains. To verify that the putative 160 open- reading frames (ORFs) of both genome isolates correspond to expressed sequences, their transcriptome maps were generated, and they showed that over 90% of the putative ORFs are transcribed. Phylogenetic analysis conducted on several genes demonstrated that the GpSGHV differs significantly from all known large dsDNA viruses of invertebrates (baculoviruses, nudiviruses, iridoviruses, ascoviruses and whispoviruses). Based on these data, and the structure of the virus particle, a new virus family named Hytrosaviridae has been accepted by the International Committee on Taxonomy of Viruses (ICTV), as proposed by the Hytrosaviridae Study Group. The name Hytrosaviridae is derived from Hytrosa, a sigla from the Greek “Hypertrophia” for hypertrophy and “sialoadenitis” for salivary gland inflammation. Two virus genera are recognized: (1) Glossinavirus , with the type species Glossina hytrosavirus and (2) Muscavirus , with the type species Musca hytrosavirus . The GpSGHV belongs to the species Glossina hytrosavirus , and the Musca domestica SGHV (MdSGHV) belongs to the species Musca hytrosavirus . The MdSGHV genome, although smaller (124,241bp) than that of the GpSGHV, produces a similar gross pathology and contains a number of putative ORFs which are homologues to those of GpSGHV and which are distinct from other insect viruses (Garcia-Maruniak et al. , 2009, J. Gen. Virol. 90: 334- 346). Further phylogenetic analyses have been performed to elucidate the relationship of hytrosaviruses to the baculoviruses and nudiviruses, as these viruses share a number of core genes involved in DNA replication, RNA transcription, and virion host cell interaction. Results showed that hytrosaviruses, nudiviruses, and baculoviruses can be considered as members of a supergroup of invertebrate-specific circular dsDNA viruses (Wang and Jehle, 2009; Wang et al. , 2010). The availability of sequence information from multiple closely related and disparate SGHVs has provided a framework to identify and design molecular probes that target common domains. These data have been used to develop universal primer pairs that detect SGHVs and generate a virus-specific PCR product from both MdSGHV and GpSGHV, whereas no non-specific PCR product is found when using genomic DNA of healthy flies as templates (Abd-Alla et al. , 2011). Therefore, these primers sets have been used to detect novel hytrosaviruses in other insects. Proteins present in the virion (proteome) of GpSGHV and of the MdSGHV were analyzed by LC-MS/MS (Kariithi et al., 2010; Maruniak-Garcia et al. , 2009). Both viruses contained an

Page 12 array of proteins that varied in their relative abundance in either the viral envelope or nucleocapsid fraction. For example, the proteome of GpSGHV contained 61 virus-encoded proteins, the most abundant proteins (GP10 and GP96) being products of the ORF10 and ORF96 genes. Significantly, these data have been used to select viral genes for the production of recombinant proteins used as antigens for the synthesis of potential neutralizing antibodies. Furthermore, these data and general information on the viral transcriptome have served as references for designing interfering RNA (RNAi) silencing technologies. The second verifiable indicator was deciphering the pathology of the SGHVs. Utilizing quantitative PCR-based diagnostic methods (Abd-Alla et al. , 2009, 2010), noninfected and infected flies have been identified and were used to demonstrate that this virus can be both vertically and horizontally transmitted in colonized tsetse flies. In addition to salivary glands that display overt pathologies, various tissues were reported to support replication (Abd-Alla et al. , 2010). Significantly, a parallel study conducted by Lietze et al. (2010) demonstrated that the MdSGHV DNA and resulting transcripts could be detected at high levels in a range of tissues without showing hypertrophy. EM studies suggested that the tracheole system extending throughout all tissues represents the alternate site of replication for SGHV. In addition to causing hypertrophy, the SGHVs also inhibit reproductive events, leading to fly sterility. Recently, it has been discovered that this virus can replicate in the corpus cardiaca; disruption in neuroendrocrine signaling at this level blocks vitellogenesis. In heterologous fly species, evidence is accumulating that the SGHVs can replicate without causing detectable hypertrophy symptoms. For example, challenge of the stable flies ( Stomoxys calcitrans) and the black dump flies ( Hydrotaea aenescens) with MdSGHV sterilized females without inducing hypertrophy (Geden et al. , 2011a, b). Replication of GpSGHV (Seibersdorf) and of MdSGHV (Florida) was attempted in 29 cell lines derived from Lepidoptera, Hymenoptera, Hemiptera, and Coleoptera. Inoculated cells were observed daily for evidence virus replication in terms of cytopathic effects and rate of cell multiplication in comparison to mock-infected cells. To date, no evidence of replication of either virus was observed in the tested cell lines, which underscores the need to generate cell lines from the tsetse fly and house fly. The third verifiable indicator was the establishment and maintenance of asymptomatic colonies. An intial step to achieve this indicator was the identification of “virus free” tsetse fly populations. In Tanzania, molecular analysis did not detect SGHV infection in colonized flies. In contrast, in Kenya, G. pallidipes and G. morsitans colonies exhibited a prevalence of ~5% symptomatic flies. These levels are comparable to those observed in the G. pallipides colony at Seibersdorf. Over the duration of the project, various participants have collected and tested field populations and have reported that the viral incidence in feral populations varies from one location to another. For example, SGHV prevalence in different wild tsetse populations from two sites in Tanzania (on the coast and inland) did not reveal infection in 200 dissected flies and in 30 flies analyzed by PCR. In Kenya, six out of eight feral populations analyzed of G. pallidipes, G. brevipalpis , and G. austeni were found to have infection rates ranging from 0-77%. In Burkina Faso, the overall prevalence of virus determined by dissection was 9%; the prevalence was 19% for G. palpalis gambiensis (Gpg), 5% for G. morsitans submorsitans (Gms), and none for G. tachinoïdes (Gt) in areas of Kénédougou, Komoé, Houet, Kossi, Sissili, and Gnangnan. The overall virus infection determined by PCR was 5%: 7% for G. palpalis gambiensis (Gpg) , 5% for G. morsitans submorsitans (Gms), and 1% for G. tachinoïdes (Gt). Symptomatic infection from the coast collected flies was 1.2% (25/2164) as compared to 0.4% (6/1725) for inland collected flies. Molecular analysis indicated a higher infection rate of 19.81% (104/525) of asymptomatic flies. Out of 525, the number of infected females were 33/216 (15.28%), and 71/309 (22.98%) males. Infection per individual tsetse species were 51/236 for G. f. fuscipes ; 11/20 for G. m. morsitans , 12/80 for G. swynnertoni and a total of 30/80 for G. pallidipes . More virus infections were recorded in male G. f. fuscipes 20.3% (48/236) than in females 1.3% (3/236). The level of infection both microscopically and by molecular analysis indicate that some species had high prevalence of the Page 13 virus than other species. G. pallidipes, G. m. morsitans and G. f. fuscipes were all infected but the level of infections was different from one species and from one location to the other. None of the G. austeni and G. brevipalpis were infected, however this could be due to the low numbers of flies analysed. A more extensive survey was conducted to investigate the prevalence and genetic diversity of GpSGHV in wild populations of G. pallidipes collected from 10 locations in 6 eastern and southern African countries. Overall, 34.08% of the analyzed flies (672/ 1,972) tested positive by nested PCR. Prevalence of GpSGHV varied widely (2 - 100%) from one geographical location to another. Generally, fly samples collected from National Parks showed higher virus prevalence than those collected elsewhere. The diversity of GpSGHV (using sequences of 5 putative viral ORFs) was found to be low. There was no correlation of the virus diversity to geographical locations. However, the GpSGHV haplotypes could be assigned to one of two distinct clades. The haplotype from the Seibersdorf colony was the most widely distributed, and was shared by forty-seven individuals in seven of the eleven locations. GpSGHV haplotypes from Ethiopian were restricted to one clade and showed the highest divergence (with 14-16 single nucleotide mutation steps) from the Seibersdorf haplotype. In conclusion, the data generated from this survey suggest that molecular- based virus management strategies could be applicable in various tsetse colonies. A fourth verifiable indicator was the development of management strategies to reduce virus loads in mass-reared tsetse flies. Antisera, generated against the intact GpSGHV virions, and various recombinant GpSGHV structural peptides (GP10, GP96, and GP1 (P74)) have been tested as bood amendments. Feeding flies with blood supplemented with these antisera significantly reduced virus levels in treated flies. These results indicate that virus-specific antisera can be used to reduce horizontal virus transmission in tsetse colonies. Similar research on the antisera targeting the MdSGHV structural peptides has demonstrated that antisera targeting the major envelope protein also neutralizes oral transmission and significantly reduces infection levels. The SGHVs code for a homologue of the p74 gene, which also is found in many insect- pathogenic DNA viruses. This gene codes for a protein that plays an important role in the initial stages of the infection by these viruses. The RNAi-based delivery technology developed for managing white spot syndrome disease in shrimp farming was adopted for managing GpSGHV. Incorporating formalin-killed E. coli expressing dsRNA directed toward p74 h into the blood meals decreased the prevalence of SGHV in the colony over two generations. However, both the instability in the plasmid construct and unpredictable expression of dsRNA precluded adoption of this control strategy. Work also has been done to investigate the impact of two antiviral drugs on viral infection in G. pallidipes and to assess their toxicity to tsetse flies. The drugs acyclovir and valacyclovir were selected for further work. G. pallidipes flies fed on blood diet with valacyclovir for three generations showed a lower mortality and higher productivity than did flies fed on blood diet that contained acyclovir. Quantitative PCR analysis assessed the effect of the antiviral drugs on viral DNA replication. The results indicate that using valacyclovir leads to a slight reduction in virus copy numbers in F0 and F1, whereas using acyclovir did not reduce the virus copy number in F0 but caused a significant reduction in virus copy number in F1. In F2, no difference in virus copy numbers was observed between the treatments and control due to a marked reduction in the control virus level. The quantitative PCR results did not show significant differences in virus copy numbers between males and females, but the results show significant reductions in virus copy numbers over generations, regardless of the treatment (including control). Some modifications were made in the antiviral drug experimental protocol to ensure a virus contamination source in the experiment. The use of valacyclovir and acyclovir in the normal colony feeding system for six months showed two interesting observations: i) flies fed on acyclovir produced few progeny, and the treatment had to be ended, while flies fed on blood supplemented with valacyclovir maintained acceptable fecundity; and ii) the virus load in the flies treated with valacylovir was not stable and slightly increased over time, which could indicate the development of virus resistance to the drug, but this observation needs to be confirmed. Fifteen additional antiviral drugs have been identified for testing.

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Several experiments were carried out to better understand virus transmission and biology. The results confirmed that symptomatic infected flies release large numbers of virus particles into the blood during feeding. Injecting virus into flies leads to increasing virus copy numbers over time, the injected female producing progeny with high virus copy number, but the injected flies did not develop hypertrophy in the same generation. After demonstrating the role of horizontal transmission through the membrane feeding system used in the tsetse flies laboratory colony, it was recommended to initiate tsetse flies with a “clean-feeding” system. One clean-feeding colony was established by feeding the fly cages first on the fresh membrane and blood and keeping these cages and their progeny fed in the same way. The remaining blood was used by the other “normal- feeding” colony. The clean-feeding colony was maintained separately from the other colony, and samples were taken regularly to assess the virus load and the prevalence of SGH. The qPCR results indicated significant decreases in the virus load in the clean-feeding colony in comparison to the normal-feeding colony. Fly dissections indicated that after three months, the clean-feeding colony became an SGH-free colony, while 10% of the normal-feeding colony still showed SGH. A fifth verifiable indicator was to demonstrate horizontal transmission of entomopathogenic fungal inoculum. Experiments conducted in Calkins and Webb’s cage to simulate field conditions have confirmed the horizontal transmission of conidia between Metarhizium anisopliae-infected G. fuscipes fuscipes and free-fungus flies when flies were allowed to pass through the fungus-treated contamination device mounted in the cage and to mate thereafter. All the fungus-treated “donor” male or female flies succumbed to fungal infection within 6.3 days with mycosis. Flies that succeeded in mating with the “donor” flies, e.g. , first- or second-line “recipients,” also died from fungal infection with mycosis within 8-12 days of mating. Fungal infection also affected the reproduction potential of flies. Female flies in the control treatment produced more puparia than did those in the fungus treatments, except for the second-line recipients, which exhibited no difference between the treatments.

3. Individual achievments during the CRP Seibersdorf, IAEA: Abd-Alla, A. Collaborators: Jehle, J., Vlak, J., Bergoin, M., Boucias, D.

Tsetse flies (Diptera: Glossinidae) are responsible for the cyclical transmission of two major diseases in sub-Saharan Africa: human African trypanosomosis (HAT or sleeping sickness), and African animal trypanosomosis (AAT or nagana). Both diseases have profound impacts on the health of livestock and humans, agricultural development and nutritional resources in sub-Saharan Africa. Due to a lack of effective vaccines and inexpensive drugs for HAT and development of drug resistance for AAT, vector control remains the most feasible strategy for the sustainable management of these diseases. The successful eradication of Glossina austeni from the Island of Unguja, United Republic of Tanzania, using an area-wide integrated pest management approach (AW-IPM) including the release of sterile male tsetse flies in (1994-1997), lead the Government of Ethiopia to embark on a program to eradicate Glossina pallidipes from the Southern Rift Valley. For this project, a laboratory colony with flies originating from the Ethiopian target areas was established at the Insect Pest Control Laboratory (IPCL) of the FAO/IAEA Agriculture & Biotechnology Laboratories, Seibersdorf, Austria. However, the productivity of this colony declined steadily over two years, leading to its collapse in 2002. The colony collapse was probably due to low productivity related to testicular degeneration (in males) and ovarian abnormalities (in females) caused by the salivary gland hypertrophy virus. Due to this frailer in colony establishment and in the frame of helping the tsetse project in Ethiopia the agency embarked a research activity aiming to identify the cause of the colony collapse and to develop management strategy to maintain the colony productivity at level reply to the SIT project need. In this frame also the agency embarked Page 15 this CRP with the aim to gather and coordinate the research groups to work together to achieve the agency goal. Considerable achievement was gained during the CRP activity. These achievements are listed below. In addition several research articles were published.

Proress during the RCP

• Development of Non-destructive PCR and qPCR specific to GpSGHV • Complete sequence and transcriptional analysis of Gp SGHV Ugandan and Ethiopian isolate • Establishment of new insect virus family Hytrosaviridae, Universal primers • Determine the virus transmission mode • Analyze the impact of the antiviral drug treatment on the virus replication and the achievement of low virus load colony using clean feedingand Valacyclovir • Optimize and analyze the impact of Antibodies, oligonucleotides, RNAi on the virus replication and infection • Complete the proteomic analysis of the virus particles • Determine the SGH prevalence and genetic variability in wild tsetse populations • Analyse the impact of SGH on tsetse mating behavior in field cage

References Abd-Alla, A., Bossin, H., Cousserans, F., Parker, A., Bergoin, M., Robinson, A., 2007. Development of a non-destructive PCR method for detection of the salivary gland hypertrophy virus (SGHV) in tsetse flies. J. Virol. Methods 139, 143-149. Abd-Alla, A.M.M., Cousserans, F., Parker, A.G., Jehle, J.A., Parker, N.J., Vlak, J.M., Robinson, A.S., Bergoin, M., 2008. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. J. Virol. 82 , 4595- 4611. Abd-Alla, A., Cousserans, F., Parker, A., Bergoin, M., Chiraz, J., Robinson, A., 2009a. Quantitative PCR analysis of the salivary gland hypertrophy virus (GpSGHV) in a laboratory colony of Glossina pallidipes . Virus Res. 139 , 48-53. Abd-Alla, A.M.M., Boucias, D.G., Bergoin, M., 2010a. Hytrosaviruses: Structure and genomic properties, in: Asgari, S., Johnson, K.N. (Eds.), Insect Virology. Caister Academic Press, Norfolk, pp. 103-121. Abd-Alla, A.M.M., Kariithi, H., Parker, A.G., Robinson, A.S., Kiflom, M., Bergoin, M., Vreysen, M.J.B., 2010b. Dynamics of the salivary gland hypertrophy virus in laboratory colonies of Glossina pallidipes (Diptera: Glossinidae). Virus Res. 150 , 103-110. Abd-Alla, A.M.M., Parker, A.G., Vreysen, M.J.B., Bergoin, M., 2011a. Tsetse Salivary Gland Hypertrophy Virus: Hope or Hindrance for Tsetse Control? PLoS Negl. Trop. Dis. 5 , e1220. Abd-Alla, A.M.M., Vlak, J.M., Bergoin, M., Maruniak, J.E., Parker, A.G., Burand, J.P., Jehle, J.A., Boucias, D.G., 2009b. Hytrosaviridae: a proposal for classification and nomenclature of a new insect virus family. Arch. Virol. 154 , 909-918. Garcia-Maruniak, A., Abd-Alla, A.M.M., Salem, T.Z., Parker, A.G., van Oers, M.M., Maruniak, J.E., Kim, W., Burand, J.P., Cousserans, F., Robinson, A.S., Vlak, J.M., Bergoin, M., Boucias, D.G., 2009. Two viruses that cause salivary gland hypertrophy in Glossina pallidipes and Musca domestica are related and form a distinct phylogenetic clade. J. Gen. Virol. 90 , 334-346. Kariithi, H.M., Ince, A.I., Boeren, S., Vervoort, J., Bergoin, M., van Oers, M.M., Abd-Alla, A., Vlak, J.M., 2010. Proteomic analysis of Glossina pallidipes Salivary Gland Hypertrophy Virus virions for immune intervention in tsetse fly colonies. J. Gen. Virol. 91 , 3065-3074.

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Montpellier 2 University, Montpellier, France: Bergoin, M. Collaborators: Francois Cousserans; Adly Abd-Alla, A., Just Vlak; Drion Boucias; Johannes Jehle Pre-period of RCP In spring of 2002, Alan Robinson sent me a letter to Max explaining the problem the Laboratory of Entomology at Seibersdorf was facing with the collapse of a tsetse colony from Ethiopia succesfully established in the late 90’s. In June of the same year I visited the Seibersdorf Laboratory and after fruitful discussions with Alan Robinson and Andrew parker it was decided that a collaborative work could be started between the Seibersdorf and the Montpellier laboratoriess to characterize the virus. Freshly dissected hypertrophied salivary glands were brought back to Montpellier, the virus was purified by sucrose density gradient, the viral DNA was extracted and a partial EcoRI –restricted DNA Library was produced. Among about a hundred of clones isolated, approximatly 12 kbp of DNA were sequenced. One of the sequences showing a strong homology with the P74 baculovirus envelope protein. Based was used to deseign specific sets of primers in order to establish a sensitive diagnostic method. In November 2002 François Cousserans brought to Seibersdorf the primers, initiated the first screening of flies and validated the PCR test. This method immediatly replaced the labour intensive and time consuming standard method of dissection and subsequent examination of salivary glands for hypertrophy. Furthermore, its high sensitivity allowed detecion the virus in asymptomatic flies. Herve Bossin joined the Entomology Unit in 2003 followed by Adly Abd-Alla in 2004 to work on SGHV. Proress during the RCP -Optimization of the PCR conditions and development of a non destructive method for virus detection based on the extraction of viral DNA from a single middle leg. Demonstration that 100% of flies of the Uganda strain of the Entomology Laboratory were infected, most of them asymptomatically. (Abd-Alla et al. 2007) -Assembly and annotation of the GpSGHV genome Uganda strain (Abd-Alla et al. 2008) -Assembly and annotation of the Ethiopian strain of GpSGHV. Demonstration that Uganda and Ethiopia strains are almost identical (to be published) -Comparison of GpSGHV and MdSGHV genome sequences and organizations (Garcia-Maruniak et al.2009, Abd-Alla, Boucias, Bergoin 2010) -Participation to the Hytrosaviridae working group of ICTV. Proposal for the classifification of MdSGHV and GpSGHV into the new family Hytrosaviriadae (Abd-Alla et al 2009b) -Improvement of diagnostic by QPCR analysis to estimate the virus loads in flies .This quantitative approach showed that the syndrome of salivary gland hyperplasia is associated with virus loads exceeding 10 8 virus copy number per fly. In addition, it allowed demonstrating that the high virus copy number rejected by symptomatic flies into the blood during feeding is the main source of virus propagation in mass rearing laboratory conditions (Abd-Alla et al. 2009a, Abd-Alla et al. 2010b, 2011). -Production of monospecific rabbit antisera against 3 immunodominant GpSGHV envelope proteins ORF 10, 47 and 96. These antisera have been used in proteomic analysis of the virus (Kariithi et al 2010 and manuscript in preparation) and in sero-neutraliztion of virions in feeding tests (Seibersdorf) -Participation in the 3D structure of GpSGHV virion by cryo-EM in collaboration with Igor Orlov and Danièle Spenher at IGBMC (Strasbourg) at identifing virion of GpSGHV Monospecific These sera the sera were used for EM immunogold labeling (Wageningen) and sero-neutralization (Seibersdorf) of virions in feeding tests. Ultrastructural analysis of the GpSGHV virion was initiated in collaboration with Dr. Danièle Sepnher and Igor Orlov, two specialists of of cryo- electron microscopy at IBMC (Strasbourg). This Institute is equipped with a sophisticated cryoEM equipment allowing the construction of 3D structures of virions at less than 10 angstrom resolution. References Page 17

Abd-Alla, A., Bossin, H., Cousserans, F., Parker, A., Bergoin, M., Robinson, A., 2007. Development of a non-destructive PCR method for detection of the salivary gland hypertrophy virus (SGHV) in tsetse flies. J. Virol. Methods 139, 143-149. Abd-Alla, A.M.M., Cousserans, F., Parker, A.G., Jehle, J.A., Parker, N.J., Vlak, J.M., Robinson, A.S., Bergoin, M., 2008. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. J. Virol. 82 , 4595- 4611. Abd-Alla, A., Cousserans, F., Parker, A., Bergoin, M., Chiraz, J., Robinson, A., 2009a. Quantitative PCR analysis of the salivary gland hypertrophy virus (GpSGHV) in a laboratory colony of Glossina pallidipes . Virus Res. 139 , 48-53. Abd-Alla, A.M.M., Boucias, D.G., Bergoin, M., 2010a. Hytrosaviruses: Structure and genomic properties, in: Asgari, S., Johnson, K.N. (Eds.), Insect Virology. Caister Academic Press, Norfolk, pp. 103-121. Abd-Alla, A.M.M., Kariithi, H., Parker, A.G., Robinson, A.S., Kiflom, M., Bergoin, M., Vreysen, M.J.B., 2010b. Dynamics of the salivary gland hypertrophy virus in laboratory colonies of Glossina pallidipes (Diptera: Glossinidae). Virus Res. 150 , 103-110. Abd-Alla, A.M.M., Parker, A.G., Vreysen, M.J.B., Bergoin, M., 2011a. Tsetse Salivary Gland Hypertrophy Virus: Hope or Hindrance for Tsetse Control? PLoS Negl. Trop. Dis. 5 , e1220. Abd-Alla, A.M.M., Vlak, J.M., Bergoin, M., Maruniak, J.E., Parker, A.G., Burand, J.P., Jehle, J.A., Boucias, D.G., 2009b. Hytrosaviridae: a proposal for classification and nomenclature of a new insect virus family. Arch. Virol. 154 , 909-918. Garcia-Maruniak, A., Abd-Alla, A.M.M., Salem, T.Z., Parker, A.G., van Oers, M.M., Maruniak, J.E., Kim, W., Burand, J.P., Cousserans, F., Robinson, A.S., Vlak, J.M., Bergoin, M., Boucias, D.G., 2009. Two viruses that cause salivary gland hypertrophy in Glossina pallidipes and Musca domestica are related and form a distinct phylogenetic clade. J. Gen. Virol. 90 , 334-346. Kariithi, H.M., Ince, A.I., Boeren, S., Vervoort, J., Bergoin, M., van Oers, M.M., Abd-Alla, A., Vlak, J.M., 2010. Proteomic analysis of Glossina pallidipes Salivary Gland Hypertrophy Virus virions for immune intervention in tsetse fly colonies. J. Gen. Virol. 91 , 3065-3074.

University of Florida, Gainesville, Florida, USA: Boucias, D. Collaborators: Abd-Alla, A., Hu. Z., and Geden, C. Our lab has examined the general pathology and impact of the Musca domestica hytrosavirus on its host to provide a framework for comparative analysis to the G. pallipides hytrosavirus. We have already reported extensively on the molecular characterization (sequence, transcriptome, and proteome) of this somewhat smaller relative of the GpSGHV. Over the final portion of this CRP we have made and published upon additional aspects of the biology of this unique group of insect viruses. • Data on the comparative pathology of the GpHV and MdSGHV viruses has in part been compiled and published (Lietze et al., 2011 J. Invertebr. Pathol. 107:161-163. • Studies on the disease dynamics and peristance of Musca domestica salivary gland hypertrophy virus has been compiled and published (Lietze et al., 2012 Appl and Environ. Microbiol. 78:311-317. • The tissue tropism of Musca domestica salivary gland hypertrophy virus (MdSGHV) infecting adult house flies was examined by transmission electron microscopy (TEM) and quantitative real-time PCR (Lietze et al., 2011, Virus Research 155:20-27. • Studies on the structural proteins from the Musca domestica hytrosavirus with an emphasis on the major envelope protein have been conducted and submitted for pubication (Boucias et al, 2012, J. Invertebr. Pathol.in press). Ongoing projects include continued work with collaborators to address the virus trafficking, potential midgut viral receptor biology. Secondly we will complete our analysis of infection on host

Page 18 neuroendrocrine relationships. Thirdly the data validating the synergistic activity of reducing agents on infection and the potential role of the peritrophic membrane in the age related resistance expressed by house flies to SGHV infection. Will be summarized and published providing a means to best formulate the virus in a house fly attractant/bait system Publications: Abd-Alla, A., D.G. Boucias, and M. Bergoin. 2010. Hytrosaviridae. Pp. 101-119 in S. Asgari and K. Johnson (Eds.), Insect Virology. Horizon Scientific Press and Caister Academic Press, Norwich, United Kingdom. Lietze, V-U., A. M. M. Abd-Alla, Vreysen, M. J.B., Geden , C. J., and D.G. Boucias . 2011. Salivary gland hypertrophy viruses (SGHVs): a novel group of insect pathogenic viruses. Ann. Rev. Entomol. 56:63–80. Lietze, V., C.J. Geden, P. Blackburn, and D.G. Boucias. 2007. Effects of Md SGHV infection on the reproductive behavior of the house fly, Musca domestica . Applied and Environmental Microbiology 73: 6811-6818. Geden, C.J., V. Lietze, and D.G. Boucias. 2008. Seasonal prevalence and transmission of salivary gland hyperplasia virus of house flies, Musca domestica L. (Diptera: Muscidae). Journal of Medical Entomology 45 (1): 42-51. Garcia-Maruniak, A., J.E. Maruniak, W. Farmerie, and D.G. Boucias. 2008. Sequence analysis of a non-classified, non-occluded DNA virus that causes salivary gland hypertrophy of Musca domestica , MdSGHV. Virology 377: 184-196. Garcia-Maruniak, A., A.M.M. Abd-Alla, T.Z. Salem, A.G. Parker, V. Lietze, M.M. van Oers, J.E. Maruniak, W. Kim, J.P. Burand, F. Cousserans, A.S. Robinson, J.M. Vlak, M. Bergoin, and D.G. Boucias. 2009. Comparative analysis of two viruses that cause salivary gland hypertrophy in Glossina pallidipes and Musca domestica . Journal of General Virology 90: 334-346. Salem, T.Z., A. Garcia-Maruniak, V.-U. Lietze, J.E. Maruniak, and D.G. Boucias. 2009. Analysis of transcripts from predicted ORFs of the Musca domestica salivary gland hypertrophy virus (MdSGHV). Journal of General Virology 90: 1270-1280. Abd-Alla, A.M.M., J.M. Vlak, M. Bergoin, J.E. Maruniak, A. Parker, J.P. Burand, J.A. Jehle, and D.G. Boucias. 2009. Hytrosaviridae : A proposal for classification and nomenclature of a new insect virus family. Archives of Virology 154: 909-918. Lietze, V.-U., K. Sims, T.Z. Salem, C.J. Geden, and D.G. Boucias. 2009. Transmission of MdSGHV among adult house flies, Musca domestica (Diptera: Muscidae), via salivary secretions and excreta. Journal of Invertebrate Pathology 101: 49-55. Prompiboon, P., V.-U. Lietze, J.S.S. Denton, C.J. Geden, T. Steenberg, and D.G. Boucias. 2010. The Musca domestica salivary gland hypertrophy virus: An insect virus that globally infects and sterilizes female house flies. Applied and Environmental Microbiology 76: 994-998. Abd-Alla, A., T. Salem, A.G. Parker, Y. Wang, J.A. Jehle, M.J.B. Vreysen, and D. Boucias. 2011. Universal primers for rapid detection of hytrosaviruses. Journal of Virological Methods 171: 280–283. Lietze,V.-U., T.Z. Salem, P. Prompiboon, and D.G. Boucias. 2011. Tissue tropism of the Musca domestica salivary gland hypertrophy virus. Virus Research 155:20-27. Lietze,V-U., Abd-Alla,A., and D.G. Boucias 2011. Two hytrosaviruses, MdSGHV and GpSGHV, induce distinct cytopathologies in their respective host insects. J Invertebr. Pathol. 107:161-163. Geden, C.J., T. Steenberg, V. Lietze, and D. Boucias. 2011. Salivary gland hypertrophy virus of house flies in Denmark: Prevalence, host range, and comparison with a Florida isolate. J. Vector Ecology 48(6): 1128-1135. Geden, C., A. Garcia-Maruniak, V.-U. Lietze, J. Maruniak, and D. G. Boucias 2011. Impact of house fly salivary gland hypertrophy virus (MdSGHV) on a heterologous host, stable fly (Stomoxys calcitrans ). J. Medical Entomology 36:231-238. Page 19

Lietze, Verena-Ulrike, Christopher J. Geden, Melissa Doyle, and Drion G. Boucias 2011. Transmission dynamics and persistence of MdSGHV in laboratory house fly ( Musca domestica ) populations. Appl. Environ. Microbiol. (accepted).

University of Massachusetts - Amherst, Amherst MA, USA: Burand, J *. Collaborators: Abd-Alla, A., Boucias, D. As with tsetse flies infected with SGHV, female H. zea moths infected with the nudivirus HzNV-2 are sterile. The sterility of these moths results from the proliferation of cells in the common and lateral oviducts and the malformation of female reproductive tissues similar to what occurs in salivary glands of infected tsetse. Both GpSGHV of the tsetse fly, G. pallidipes, and HzNV-2 contain an ortholog of the p74 gene which codes for a virus structural protein involved in entry into cells, the first step in virus infection of the host. We have demonstrated that dsRNA containing sequences homologous to either HzNV-2 p74 gene or the nonstructural protein gene coded by ORF7, blocks viral pathology in infected female moths. These results suggest that dsRNAs complimentary to viral sequences function as RNAi and down-regulate expression of these genes, ultimately blocking virus replication and transmission.

Wuhan Institute of Virology, Chinese Academy of Sciences: Zhihong Hu Collaborators: Drion G. Boucias, Just M.Vlak The objective of our research is to generate antibodies that could be used for other members of the CRP for pathology study of hytrosavirus, as well as to study if the antibodies could suppress the virus infection in flies. Orf22 , orf86 , orf96 and iap genes from MdSGHV, and pif1 , pif2 , orf140 and orf261R from GpSGHV were successfully expressed in E. coli and polyclonal antibodies were generated from rabbits. Western blot analyses showed that anti-MdSGHV22, anti-MdSGHV86 and anti-MdSGHV96 could detect MdSGHV virus and virus infected gland. Immune-gold electron microscopy showed that anti-MdSGHV86 reacted to the nucleocapsid of MdSGHV, where anti- MdSGHV96 sera recognized target antigens associated with the virus envelope, confirmed that MdSGHV96 is the major envelope protein of virus. Oral bioassays demonstrated that anti- MbSGHV96 polyclonal sera could suppress the level of orally acquired infections of MdSGHV. Therefore, the objectives have been accomplished. • A serial of polyclonal antibodies against structural proteins of MdSGHV and GpSGHV were generated. • Antibodies against MdSGHV ORF86 and ORF96 could detect viral nucleocapsid and envelope respectively, therefore can be used for pathology study. • Bioassay showed that anti-MdSGHV96 could suppress the level of orally acquired infections of MdSGHV. Publication: D. G. Boucias, F. Deng, Z. Hu, A. Garcia-Maruniak, and V.-U. Lietze. 2012. Analysis of the Structural proteins from the Musca domestica hytrosavirus with an emphasis on the major envelope protein Journal of Invertebrate Pathology. In press .

JKI - Institute for Biological Control. Darmstadt, Germany: Jehle, J. Collaborators: Abd-Alla, A., Bergoin, M., Boucias, D., Burand, J.,Vlak, J, Based on the complete genome sequence of GpSGHV and the MdSGHV twelve open reading frames (ORFs) were identified in both genomes that are homologous to baculovirus and nudivirus core genes. These include genes encoding subunits of the RNA polymerase (lef-4, lef-5, lef-8, lef- 9), genes involved in DNA replication (DNApol, hel), genes important for virion host cell interaction (p74, pif-1, pif-2, pif-3) and genes of unknown function (p33, ac81). In addition, several other baculovirus and nudivirus homologous not belonging to the core gene set of these viruses are

Page 20 present in the genomes of SGHVs, e.g. a gene homologue for polh/gran , the baculovirus occlusion body protein was found in the genome of MdSGHV. Though different phylogenetic analyses have produced some conflicting results, it is proposed that SHGVs, baculoviruses and nudiviruses share a common ancestor and are related phylogenetically. A new virus family Hytrosaviridae consisting of the two genera Glossinavirus and Muscavirus has been established and was approved by the International Committee on Taxonomy of Viruses (ICTV). • In silico sequence and database comparison of the SGHV genes with selected gene sequences from nudiviruses and baculoviruses. • Assisting in the in phylogenetic analyses of selected SGHV genes. • Design of degenerate oligonucleotides for PCR detection of GpSGHV • Assisting in drafting a taxonomic proposition for SGHV

Publications: JEHLE , J.A., WANG , Y. ABD ALLA , A. M. M. (2012). Evolution, Phylogeny and Taxonomy of Hytrosaviridae (Minireview, submitted to Journal of Invertebrate Pathology ) WANG , Y., BININDA -EMONDS , O. R. P., VAN OERS , M. M., VLAK , J. M., JEHLE , J. A. (2011). Nudiviruses give insights into the evolution of nuclear arthropod-specific large circular double- stranded DNA viruses. Virus Genes 42 , 444-456). WANG , Y., BININDA -EMONDS , O. R. P., JEHLE , J. A. (2012). Nudivirus Genomics and Phylogeny. In: Viral Genomes - Molecular Structure, Diversity, Gene Expression Mechanisms and Host-Virus Interactions, Maria Laura Garcia and Victor Romanowski (Ed.), ISBN: 978-953-51-0098-0, InTech, Available from: http://www.intechopen.com/articles/show/title/nudivirus-genomics-and- phylogeny ABD -ALLA , A. M. M., SALEM , T. Z., PARKER , A. G., WANG , Y., JEHLE , J. A., VREYSEN , M. J. B., BOUCIAS , D. (2011). Universal primers for rapid detection of hytrosaviruses. Journal of Virological Methods 171 , 280-283. WANG , Y., JEHLE , J. A. (2009). Nudiviruses and other large, double-stranded circular DNA viruses of invertebrates: New insights on an old topic. Journal of Invertebrate Pathology 101 , 187-193. ABD -ALLA , A. M. M., VLAK , J. M., BERGOIN , M., MARUNIAK , J. E., PARKER , A. G. BURAND , J. P., JEHLE , J. A., BOUCIAS D. G (2009). Hytrosaviridae : a proposal for classification and nomenclature of a new insect virus family. Archives of Virology 154 , 909-918. ABD -ALLA , A. M. M., COUSSERANS , F., PARKER , A., JEHLE , J. A., PARKER , N., VLAK , J. M., ROBINSON , A., BERGOIN , M. (2008). Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. Journal of Virology 82 , 4595-4611.

Wageningen University, The Netherlands: Vlak J.M., Collaborators: H. Karrithi, Bergoin, M., Koekemoer, O., Abd-Alla, A., Jehle, J., Hu, ZH, Aksoy, S.

The repertoire of proteins comprising intact virions of Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) is likely to have significant consequences for virus morphology and morphogenesis. In this study, cryo-tomography of GpSGHV from hypertrophied salivary glands (SGs) of Glossina pallidipes revealed that the virion envelope and nucleocapsid components are separated by a jacket of proteinaceous material. We concluded that this material represents the GpSGHV tegument. The entire lengths of mature, extracellular virions consisted of well-organized spiral-like ridges containing regular proteinaceous substructures. Further, while nucleocapsids were restricted to the nuclei, enveloped virions were essentially located in the cell cytoplasm. This result, coupled to observation of nucleocapsids passing out of the nucleus via the nucleopore, suggests that GpSGHV nucleocapsids are assembled in the nucleus and egress to the cytoplasm of infected SG cell where virion envelopment is Page 21 orchestrated. It seems that the entire envelopment occurs in the cytoplasm of infected cells and that the virions are released into the salivary gland by cell rupture or disintegration. Since the GpSGHV virions were highly unstable in solution, we have amended the original purification procedure (Kariithi et al., 2012) and arrived at a highly purified preparation of intact virions as evidence by electron microscopy. These virions were fractionated into nucleocapsids and envelopes by two rounds of 1% Nonidet P-40-treatment, followed by 12% SDS-PAGE separation and subsequent protein identification by tandem mass spectrometry (LC-MS/MS) or defined antibodies. Fifty-four distinct, virally-encoded proteins were reliably identified by ≥ 2 peptides per protein, of which at least one was unique. We assigned 10 of these proteins to the envelope, 15 to the nucleocapsid and 29 to the tegument. In contrast to what has been found previously (Kariithi et al. , 2010) the product of orf10 , a 130 kDa protein is now found in the tegument and not in the envelope. As expected the peroral infectivity factors (PIFs) were exclusively found in the membrane fraction of the virion. Analyses of posttranslational modifications revealed that that several of the virion proteins are serine- /threonine-phosphorylated and that the proteins encoded by ORFs 38, 39, 40, 62 and 97 are O-glycosylated. Additionally, 56 virion-associated (cellular) proteins were identified, of which the presence of α-tubulin, γ-actin and ubiquitin was verified by a proteinase K protection assay and immunoblotting. Bioinformatics analyses of the identified virion-associated GpSGHV protein sequences revealed motifs with functional and structural implications on the virus morphogenesis. Among these is the involvement of the endoplasmic reticulum and Golgi in the morphogenesis of the virus, which is compatible with the cytoplasmic assembly of GpSGHV. Our findings provide new insights into GpSGHV morphology and morphogenesis, and point us towards areas of the virus pathogenesis in our future studies. Furthermore, the proteomic analysis may call for re-evaluation of the immuno-intervention strategy to control GpSGHV infection in tsetse fly rearing facilities for the sterile insect technique by using the newly identified envelope proteins as a basis for antibody development and GpSGHV intervention studies (Kariithi et al., 2010). The above research were the result of a previous effort in determining the structure and sequence of the GpSGHV genome (Abd-Alla et al., 2008) and the comparison with the related hytrosavirus, Musca domestica SGHV (MdSGHV) (Garcia-Maruniak et al., 2009). This not only allowed the identification and gene assignment of virion proteins on the viral genome, but also the establishment of a new virus family Hytrosaviridae (Abd-Alla et al, 2009), which is now accepted by the Inernational Committee on Taxonomy of Viruses. To successfully execute virus intervention studies it was necessary to study the genetic diversity of GpSGHV in African tsetse flies. From these studies it was concluded that the genetic variation was minimal (less than 1%). This implies that the virus intervention studies using antibodies are in principle generally applicable in mass rearings of tsetse. The general conclusion is that immune intervention to block transmission of GpSGHV is generally applicable, but requires fine-tuning and testing beyond the laboratory scale.

References Abd-Alla, A., F. Cousserans, A. Parker, J.A. Jehle, N. Parker, J.M. Vlak, A. Robinson and M. Bergoin. 2008. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. Journal of Virology 82: 4595-4611 Garcia-Maruniak, A., A.M.M. Abd-Alla, T.Z. Salem, A.G. Parker, V-U. Lietze, M.M. van Oers, J.E. Maruniak, W. Kim, J.P. Burand, F. Cousserans, A.S. Robinson, J.M. Vlak, M. Bergoin and D.G. Boucias. 2009. Two viruses that cause salivary gland hypertrophy in Glossina pallidipes

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and Musca domestica are closely related and form a distinct phylogenetic clade. Journal of General Virology 90: 334 - 346. Abd-Alla, A M.M., J.M. Vlak, M. Bergoin, J. Maruniak, A. Parker, J.P. Burand, J.A. Jehle and D.G. Boucias. 2009. Hytrosaviridae : a proposal for classification and nomenclature of a new insect virus family. Archives of Virology 154: 909-918. Henry M. Kariithi, H.M., , I.A. Ince, S. Boeren, J. Vervoort, M. Bergoin, M.M. van Oers, A.M.M. Abd-Alla and J.M. Vlak. 2010. Proteomic analysis of Glossina pallidipes Salivary Gland Hypertrophy Virus virions for immune intervention in tsetse fly colonies. Journal of General Virology 91: 2985-2993. Kariithi, H.M., I.A. Ince, S.A. Boeren, A.M.M. Abd-Alla, Andrew G. Parker, J.M. Vlak, S. Aksoy, J.M. Vlak and M.M. van Oers. 2011. The salivary secretome of the tseste fly Glossina pallipides (Diptera: Glossinidae) infected by salivary gland hypertrophy virus. PLoS Neglected Tropical Diseases 5(11): e1371. Kariithi, H.M., M. Ahmadi, A.G. Parker, A.S. Robinson, G. Franz, V. Ros, I. Haq, A.M. Elashry, J.M. Vlak, M. Bergoin, J.B. Vreijsen and A.M.M. Abd-Alla. 2012. Prevalence and genetic variation of salivary gland hypertrophy virus in the wild populations of the tsetse fly Glossina pallipides (Diptera: Glossinidae) from southern and eastern Africa. Journal of Invertebrate Pathology, to be revised. Abd-Alla, A.M.M., M. Bergoin, A.G. Parker, N.K. Maniania, J.M. Vlak, K. Bourtzis, D.G. Boucias and S. Aksoy. 2012. Improving SIT for tsetse flies through research on their symbionts and pathogens. Journal of Invertebrate Pathology, to be revised. Kariithi, H.M., I.A. Ince, M.M. van Oers, A.M.M. Abd-Alla and J.M. Vlak. 2012. Proteomic footprints of a member of Glossinavirus (Hytrosaviridae ): An expeditious approach to virus control strategies in tsetse factories. Journal of Invertebrate Pathology, to be revised.

Great Lakes Forestry Centre, Canada: Arif, B., Collaborators: Abd-Alla, A., Geden, C. S., Hu, Z.

Over the past three years, attempts to replicate the GSGHV and MSGHV in cell lines existing at the Great Lakes Forestry Centre were conducted. Of the collection of approximately 129 cell lines stored in liquid nitrogen at the Centre, 29 derived from Lepidoptera, Hymenoptera, Coleoptra and Hempitera were selected to test the replication of the two viruses. The selected lines originated mostly from embryos or ovaries but also from other tissues such as the midgut. Each line was seeded into 48 flat well plates, allowed to attach then inoculated with either GSGHV of MSGHV and let adsorb for 1 hour. Medium (100 ul) was added to each well and the cells were examined twice daily for evidence of virus replication in comparison to mock infected cells. No evidence of virus replication was observed in the inoculated cells in terms of cells’ doubling time, cytopathic effects, occasional EM observations or amplification by PCR of a sequence in the MSGHV genome. Both the mock infected and inoculated cells appear to grow at approximately the same rate.

USDA, ARS, CMACE, Gainesville, Florida, USA: Geden, C. Collaborators: Boucias, D., Maniania, N. K., Abd-Alla, A. Accomplishments: • Conducted field surveys to determine seasonal prevalence of MdSGHV in US house flies; identified strong correlation between prevalence and fly density; ruled out vertical transmission; documented horizontal transmission via contaminated food, water, and cages (Geden et al. 2008b) • Identified promising attractant devices that can be modified to deliver MdSGHV (Geden et al. 2008a, 2009, 2012). Page 23

• Conducted field survey of house flies in Denmark for MdSGHV; obtained several Danish virus isolates and compared virulence of Florida and Danish isolates; observed no SGH in 6 species of flies that occur sympatrically with house flies; conducted host range studies and observed reproductive suppression in virus-injected stable fly ( Stomoxys calcitrans ) and black dump fly (Hydrotaea aenescens ) (Geden et al 2011a). • Observed MdSGHV replication in stable fly fat body, ovaries and salivary glands; documented virus loads required to shut down ovarian development in this host; noted absence of SGH symptoms in infected stable flies (Geden et al. 2011b) • Evaluated various dry and liquid bait vehicles for per os transmission of MdSGHV to house flies; found that application of viral homogenates to fly resting sites or directly on flies resulted in substantially higher transmission rates than virus-treated food baits (Geden et al. 2011a; Geden et al 2012, in prep).

Publications: Geden, C. J. 2008a. Visual targets for capture and management of house flies, Musca domestica . J. Vector Ecol. 31: 152-157. Geden C. J., V. Lietze, and D.G. Boucias. 2008b. Seasonal prevalence and transmission of salivary gland hypertrophy virus of house flies (Diptera:Muscidae). J. Med. Entomol. 45: 42-51. Geden, C. J., D. E. Szumlas and T.W. Walker. 2009. Evaluation of commercial and field- expedient baited traps for house flies, Musca domestica L. (Diptera: Muscidae). J. Vector Ecol. 34(1): 99-103. Geden, C. J., A. G. Maruniak, V.-U. Lietze, J. Maruniak, and D. G. Boucias. 2011a. Impact of house fly salivary gland hypertrophy virus (MdSGHV) on a heterologous host, stable Fly (Stomoxys calcitrans ). J. Med. Entomol. 48: 1128-1135 Geden, C. J., T. Steenberg, V.-U. Lietze, and D. G. Boucias. 2011b. Salivary gland hypertrophy virus of house flies in Denmark: Prevalence, host range, and comparison with a Florida isolate. J. Vector Ecol. 36: 231-238. Geden, C. J. and G. G. Devine. 2012. Pyriproxyfen and house flies (Diptera: Muscidae): effects of direct exposure and use of flies as autodissemination vehicles. J. Med. Entomol.

KARI-Trypanosomiasis Research Centre (TRC), Muguga, Kenya: Wamwiri, F. Trypanosome and SGHV co-infection in G.pallidipes and G.austeni in Kenya Collaborators: Robert Changasi and Sam Guya

The salivary gland virus that causes hypertrophy and reproductive abnormalities in tsetse flies has been reported from G.austeni , G.pallidipes and G.brevipalpis from nine populations in Kenya. A total of 773 flies comprising 414 G.pallidipes , 269 G.austeni and 90 G.brevipalpis were analysed. Infection prevalence obtained were 41.9%, 72.9% and 18.6% respectively. We have previously evaluated the co-infection levels of SGHV with the symbionts Wolbachia and Sodalis, and found no distinct correlation between them in G.austeni at the Kenyan coast. However, the interaction between virus and trypanosome infection is not clear. Therefore, the objective of the current study was to evaluate the interaction between trypanosome and virus infection in both naturally and experimentally infected populations. Natural infection was evaluated from sympatric G.pallidipes and G.austeni collected from the coastal belt. Experimental infection was assessed by in vivo infection of teneral G.pallidipes males using the isolates TBR KETRI3738, TBB KETRI3386 and TC KETRI3805. The flies were maintained for 35, 35 and 29 days post infection respectively after which PCR was performed using universal ITS1 primers for trypanosome detection and Gp SGHV2 primers for virus detection. Results indicate virus infection rates of 18.4% and 36.4% for G.pallidipes (N=141) and G.austeni (N=182) respectively, while trypanosome infections rates were 16.3% and 19.2% respectively. The proportion of trypanosome infected flies was higher in virus-

Page 24 negative than in virus-positive flies, however this was not significant. In the experimentally-infected flies, infection rates of 70.3%, 10% and 10% respectively were established. All flies were SGHV- negative despite the colony flies having a global infection of about 13%. These preliminary results point to virus infection having a negative correlation with trypanosome infection. Future studies will focus further on infection studies and will attempt experimental clearing and re-infection with virus innoculum.

TTRI, Tanga, Tanzania: Malele, I. Collaborators: K. Bourtzis, J. Maniania, J. van den Abbeele and A. Abd-Alla, The primary objective was to conduct surveillance of SGHV in wild and lab flies both microscopically and by using molecular methods. The target species were G. pallidipes and G. brevipalpis, however in addition, other species which include G. swynnertoni, G. f. fuscipes and G. austeni were analysed. Flies were trapped from two study sites which include the site along the coast where we trapped G. pallidipes, G.m.morsitans, G. brevipalpis and G. austeni . From the inland, flies trapped were G. pallidipes, G. swynnertoni and G. f. fuscipes . Microscopic screening of 184 lab flies of different ages were all negative. Symptomatic infection from the coast collected flies was 1.2% (25/2164) as compared to 0.4% (6/1725) for inland collected flies. Molecular analysis indicated a higher infection rate of 19.81% (104/525) of asymptomatic flies. Out of 525, the number of infected females were 33/216 (15.28%), and 71/309 (22.98%) males. Infection per individual tsetse species were 51/236 for G. f. fuscipes ; 11/20 for G. m. morsitans , 12/80 for G. swynnertoni and a total of 30/80 for G. pallidipes . More virus infections were recorded in male G. f. fuscipes 20.3% (48/236) than in females 1.3% (3/236). The level of infection both microscopically and by molecular analysis indicated that some species had high prevalence of the virus than other species. G. pallidipes, G. m. morsitans and G. f. fuscipes were all infected but the level of infections was different from one species and from one location to the other. None of the G. austeni and G. brevipalpis were infected, however this could be due to the low numbers of flies analysed. Lessons learnt from this work indicate that the virus in wild tsetse populations is not species, sex, age and season dependent but levels of infection differs from one species to the other.

Studies on tsetse symbionts ( Wolbachia and Sodalis ) indicated that the occurrence of Wolbachia in wild tsetse was very common. Three types of primers were used which include Wsp, 16S RNA and FbPA . The primers detected the Wolbachia from the three species although levels differed from one species to the other. Prevalence of Wolbachia infection in both species was common and we recorded an infection of 13/18 in G. m. morsitans and (11/18) in G. pallidipes a sympatric species. Wolbachia infection was detected in colonized G. pallidipes but not in G. brevipalpis. The infection was 10/30 by Wolbachia wsp gene primers and 8/30 by the 16S rRNA gene primers. Infection with Sodalis was also investigated for the wild tsetse flies. The infections were common in the two species analysed. The infections were 12/20 in G. pallidipes and 16/18 in G.m.morsitans .

Symptomatic infection from the coast collected flies was 1.2% (25/2164) as compared to 0.4% (6/1725) for inland collected flies. Molecular analysis indicated a higher infection rate of 19.81% (104/525) of asymptomatic flies. Out of 525, the number of infected females were 33/216 (15.28%), and 71/309 (22.98%) males. Infection per individual tsetse species were 51/236 for G. f. fuscipes ; 11/20 for G. m. morsitans , 12/80 for G. swynnertoni and a total of 30/80 for G. pallidipes . More virus infections were recorded in male G. f. fuscipes 20.3% (48/236) than in females 1.3% (3/236). The level of infection both microscopically and by molecular analysis indicates that some species had high prevalence of the virus than other species. G. pallidipes, G. m. morsitans and G. f. fuscipes were all infected but the level of infections was different from one species and from one location to the other. None of the G. austeni and G. brevipalpis was infected, however this could be due to the low numbers of flies analysed.

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Laboratoire National D’Elevage: Gisèle M.S.Ouédraogo / Sanon Collaborators: Issa Sidibé, Anicet Georges Ouédraogo, Adly M.M. Abd-Alla, Charles Eloge Lamien, Max Bergoin, J. Van Den Abbeele.

In total 1433 flies from four species ( Glossina palpalis gambiensis, Glossina tachinoides, Glossina morsitans submorsitans and Glossina medicorum) were caught, of which 296 were dissected and 544 used for screening. Out of 296 flies dissected, all species inclusive, 23 flies had symptomatic GpSGHV, corresponding to 7.8%. Glossina palpalis gambiensis had the highest percentage of infection of 14. 9%. Twenty-two (22%) of 544 flies (118 flies), all species inclusive, were infected when PCR was performed. The prevalence of infetion was highest (37%) in G. palpalis gambiensis. Among the localities, Houet was the most infected with a prevalence of 59.4%.When the gene P74 Nes 2F was used, the analysis of samples with the Seibersdorf samples show that one sequence from Glossina tachinoides had two SNPs G91/A and A337/G.

ICIPE, Kenya: Maniania, N.K. Collaborators: A. Abd-Alla, Boucias, D., F. Wamwiri, I. Malele, P. Takac, C. Geden Use of fungi for tsetse control (Maniania)

Studies have shown that tsetse flies are susceptible to fungal infection in the laboratory; but only one study has shown their use in tsetse suppression in field conditions. Fungal conidia are released in the field through a contamination device (Cd) that attracts flies that come into the trap where they get contaminated with spores before they return to the environment where they may contaminate other flies during mating. Fundamental to this approach is the efficient horizontal transmission of the pathogen to susceptible individuals within the insect population. Moreover, fungal infection can raise behavior changes in the host, some of which may compromise horizontal transmission of the inoculum. Horizontal transmission of entomopathogenic fungi in tsetse flies is well documented in the laboratory. However, there are no data to support this in the field. The objective of this component of the project was, therefore, to (i) investigate the transmission of fungal conidia between fungus-infected flies and healthy flies under field-cage conditions, (ii) the effect of fungal infection on some of the behavioral changes in tsetse flies.

1. Experiments conducted in Calkins and Webb’s cage to simulate field conditions have confirmed horizontal transmission of conidia between Metarhizium anisopliae-infected G. fuscipes/G. pallidipes and healthy flies when flies were allowed to pass through the fungus-treated contamination device mounted in the cage and to mate thereafter. All the fungus-treated “donor” male or female flies succumbed to fungal infection with mycosis. Most of flies that succeeded to mate with the “donor” flies, first or second line “recipient” also died from fungal infection with mycosis. Generally, mortality was faster among “donor” flies, followed by first line “recipient” and lastly by second line “recipient”. This can be explained by the amount of inoculum picked by flies. For instance, single “donor” fly could pick as much as 3.5 x 10 6 conidia while first and second line “recipient’ flies will pick 5.0 x 10 5 and 1.2 x 10 4 conidia/fly by, respectively.

2. We have also demonstrated the inoculum transmission from flies emerging from pupae. G. fuscipes and G. pallidipes flies emerging from pupae buried previously in fungus-contaminated sand were able to pick up fungal infection and transferred fatal inoculum to “recipient” flies during mating.

3. Fungus-infected irradiated male flies were also able to transfer inoculum to healthy flies before they died although the rate of fungal infection was generally low compared to other previous studies on horizontal transmission. No plausible explanation for these results. Nevertheless,

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there was no significant difference in the number of female flies mating with irradiated male “donor” between the control and the fungus-treated ones.

4. Fungal infection did not have effect on the mating behavior such as competitiveness, duration of copulation and the number of jerking.

5. However, fungal infection reduced blood meal intake and reproduction potential of flies. Fungus-infected female flies produced fewer pupae than the fungus-free ones.

6. Publication: Maniania N.K. and Ekesi S. The use of entomopathogenic fungi in the control of tsetse flies. Submitted to Journal of Invertebrate Pathology.

Yale University, USA: Aksoy, S. Collaborators: Heddi, A., Malacrida, A., Bourtzis, K., Wolfgang, M and Takac, P.

Abstract : During this CRP program, we worked on the role of tsetse symbionts Wigglesworthia and Wolbachia on host fecundity and reproduction and the impact of tsetse symbionts on host immunity. Although the presence of symbiotic microbes, especially Wigglesworthia was shown to be essential for fecundity, the molecular basis of its symbiotic role remained unknown. Similarly, although laboratory and natural populations of tsetse have been shown to harbour Wolbachia , the functional role of Wolbachia in tsetse biology remained unknown. We showed that loss of fecundity due to Wigglesworthia can be rescued by supplementing the host diet with yeast. We developed and used the tsetse lines that are free of Wigglesworthia and all symbionts, respectively to investigate host-symbiont interactions. Our results showed that absence of Wigglesworthia from larval development adversely impacts cellular immunity in emerging adults and makes tsetse unusually susceptible to trypanosome infections. Our results also showed that Wolbachia infections confer strong CI in tsetse and mathematical modelling support the use of CI to derive genetically modified symbionts to replace susceptible populations in nature. Finally, we investigated the density dynamics of symbionts and SGHV in lines that lack Wigglesworthia only or that are aposymbiotic. Our results show that the tsetse host strongly regulates the density dynamics of its microbiome and that absence of Wigglesworthia compromises Sodalis fitness and that SGHV is negatively impacted by host immune responses elicited against the symbionts, especially Wolbachia . We have also collaborated with other members of this CRP on, Wolbachia infection prevalence in natural tsetse populations (DOUDOUMIS et al. 2011), presence of polyandry in tsetse (BONOMI et al. 2011) and on the molecular aspects of SGHV proteome (KARIITHI et al. 2011).

Introduction: Tsetse flies have a highly regulated and defined microbial fauna made of 3 bacterial symbionts (obligate Wigglesworthia glossinidia , commensal Sodalis glossinidius and parasitic Wolbachia pipientis) in addition to a DNA virus ( Glossina pallidipes Salivary gland Hypertrophy Virus, GpSGHV). The microbiomes are vertically transmitted from mother to offspring during this insect’s unique viviparous mode of reproduction. Many animals including insects such as mosquitoes rely on the presence of symbiotic bacteria for proper immune system function. However, the molecular mechanisms that underlie this phenomenon are poorly understood including their role in tsetse. We summarized the state of this knowledge in a review paper (WEISS and A KSOY 2011) Similarly, although the presence of Wolbachia has been found in tsetse, its role on tsetse reproduction is unknown. This is largely because elimination of tsetse’s symbiont Wigglesworthia renders tsetse sterile preventing the development of lines where these functions can be experimentally tested. Finally, we hypothesized that there is a strong community influence that determines the population Page 27 dynamics of individual symbionts as well as SGHV. We describe below briefly the experiments performed that allowed us to investigate these interactions.

1. Development of lines that lack tsetse symbionts . We investigated the transmission route of tsetse’s endosymbionts which showed the presence of two populations of Wigglesworthia , an intracellular state in the bacteriome and an extracellular state in the milk (ATTARDO et al. 2008). This knowledge allowed us to develop lines that are cured of their Wigglesworthia infections (PAIS et al. 2008).

2. Role of symbionts on host immunity . We characterized the immune phenotype of tsetse that developed in the absence of all of their endogenous symbiotic microbes. Larval tsetse that undergo intrauterine development in the absence of their obligate mutualist, Wigglesworthia , exhibit a compromised immune system during adulthood (WEISS et al. 2011). We found that aposymbiotic tsetse ( Gmm Apo ) present a severely compromised immune system that is characterized by the absence of phagocytic hemocytes and atypical expression of immunity-related genes. The susceptible phenotype exhibited by Gmm Apo adults can be reversed when they receive hemocytes transplanted from wild-type donor flies prior to infection (WEISS et al. 2012). Furthermore, the process of immune system development can be restored in intrauterine Gmm Apo larvae when their moms are fed a diet supplemented with Wigglesworthia cell extracts. Our finding that molecular components of Wigglesworthia exhibit immuno-stimulatory activity within tsetse is representative of a novel evolutionary adaptation that steadfastly links an obligate symbiont with it’s host.

3. Role of Wolbachia in tsetse biology . Infections with the parasitic bacterium Wolbachia are widespread in insects and cause a number of reproductive modifications, including cytoplasmic incompatibility (CI). There is growing interest in Wolbachia, as CI may be able to drive desired phenotypes such as disease resistance traits, into natural populations. Although Wolbachia infections had been reported in tsetse, their functional role was unknown. This is because attempts to cure tsetse of Wolbachia by antibiotic treatment damages the obligate mutualist Wigglesworthia , without which the flies are sterile. Using the Wolbachia free and still fertile tsetse lines, we performed mating experiments for the first time, which provides evidence of strong CI in tsetse. We have incorporated our empirical data in a mathematical model and show that Wolbachia infections can be harnessed in tsetse to drive desirable phenotypes into natural populations in few generations. This finding provides additional support for the application of genetic approaches, which aim to spread parasite resistance traits in natural populations as a novel disease control method. Alternatively, releasing Wolbachia infected males can enhance Sterile Insect applications, as this will reduce the fecundity of natural females either uninfected or carrying a different strain of Wolbachia (ALAM et al. 2011).

4. Community dynamics regulating symbiotic densities. It has been possible to rear flies in the absence of either Wigglesworthia or in totally aposymbiotic state by dietary supplementation of tsetse’s bloodmeal. In the absence of Wigglesworthia , tsetse females are sterile, and adult progeny are immune compromised. The functional contributions for Sodalis are less known, while Wolbachia cause reproductive manupulations known as Cytoplasmic Incompatibility (CI). High GpSGHV virus titers result in reduced fecundity and lifespan, and have compromised efforts to colonize flies in the insectary for large rearing purposes. We investigated the within community effects on the density regulation of the individual microbiome partners in tsetse lines with different symbiotic compositions. We show that absence of Wigglesworthia results in loss of Sodalis in subsequent generations possibly due to nutritional dependancies between the symbiotic partners. While an initial decrease in Wolbachia and GpSGHV levels are also noted in the absence of Wigglesworthia , these infections eventually reach homeostatic levels indicating adaptations to the new host immune environment or nutritional ecology. Absence of all bacterial symbionts also results in an initial reduction of viral titers, which recover in the second generation. Our findings

Page 28 suggest that in addition to the host immune system, interdependencies between symbiotic partners result in a highly tuned density regulation for tsetse’s microbiome.

Publications resulting from this CRP ALAM , U., J. M EDLOCK , C. B RELSFOARD , R. P AIS , C. L OHS et al. , 2011 Wolbachia Symbiont Infections Induce Strong Cytoplasmic Incompatibility in the Tsetse Fly Glossina morsitans. PLoS Pathog 7: e1002415. ATTARDO , G. M., C. L OHS , A. H EDDI , U. H. A LAM , S. Y ILDIRIM et al. , 2008 Analysis of milk gland structure and function in Glossina morsitans: milk protein production, symbiont populations and fecundity. J Insect Physiol 54: 1236-1242. BONOMI , A., F. B ASSETTI , P. G ABRIELI , J. B EADELL , M. F ALCHETTO et al. , 2011 Polyandry is a common event in wild populations of the Tsetse fly Glossina fuscipes fuscipes and may impact population reduction measures. PLoS Negl Trop Dis 5: e1190. DOUDOUMIS , V., G. T SIAMIS , F. W AMWIRI , C. B RELSFOARD , U. A LAM et al. , 2011 Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina ). BMC Microbiology 12 (Suppl 1): S3. KARIITHI , H. M., I. A. I NCE , S. B OEREN , A. M. A BD -ALLA , A. G. P ARKER et al. , 2011 The Salivary Secretome of the Tsetse Fly Glossina pallidipes (Diptera: Glossinidae) Infected by Salivary Gland Hypertrophy Virus. PLoS Negl Trop Dis 5: e1371. PAIS , R., C. L OHS , Y. W U, J. W ANG and S. A KSOY , 2008 The obligate mutualist Wigglesworthia glossinidia influences reproduction, digestion, and immunity processes of its host, the tsetse fly. Appl Environ Microbiol 74: 5965-5974. Wang, J., Brelsfoard, C. Wu, Y. and Serap Aksoy , Intercommunity effects on microbiome and GpSGHV density regulation in tsetse flies, JIP Tsetse Symposium, under review WEISS , B., and S. A KSOY , 2011 Microbiome influences on insect host vector competence. Trends Parasitol. WEISS , B., M. M ALTZ and S. A KSOY , 2012 Obligate symbionts activate immune system development in the tsetse fly. Journal of Immunology. WEISS , B. L., J. W ANG and S. A KSOY , 2011 Tsetse immune system maturation requires the presence of obligate symbionts in larvae. PLoS biology 9: e1000619.

Institut National des Sciences Appliquées de Lyon (INSA Lyon): Abdelaziz Heddi Collaborators: Serap Aksoy, Yale University, USA Abstract During this CRP program, we have studied the precise tissue distribution of tsetse endosymbiotic bacteria and have infered their respective way of transmission from parents to progenies. To this end, we conducted a Fluorescence In situ Hybridization (FISH) study to survey bacterial spatial distribution across the fly tissues. We showed that bacteriocytes are mono-infected with Wigglesworthia , while both Wigglesworthia and Sodalis are present in the milk gland lumen. Sodalis was further seen in the uterus, spermatheca, fat body, milk and intracellular in the milk gland cells. Contrary to Wigglesworthia and Sodalis, Wolbachia were the only bacteria infecting oocytes, trophocytes, and embryos at early embryonic stages. Furthermore, Wolbachia were not seen in the milk gland and in the fat body. This work further highlights the diversity of symbiont interactions in multipartner associations and supports two maternal routes of symbiont inheritance in the tsetse fly: Wolbachia through oocytes, and, Wigglesworthia and Sodalis by means of milk gland bacterial infection at early post-embryonic stages.

Introduction The tsetse fly Glossina is the vector of the protozoan Trypanosoma brucei spp. that causes Human and Animal African Trypanosomiasis in sub-Saharan African countries. To supplement their unbalanced vertebrate bloodmeal diet, flies permanently harbor the obligate bacterium Wigglesworthia glossinidia, which resides in bacteriocytes in the midgut bacteriome organ as well Page 29 as in milk gland organ. Tsetse flies also harbor the secondary facultative endosymbionts (S- symbiont) Sodalis glossinidius that infects various tissues and Wolbachia that infects germ cells. However, while the tsetse symbiont functions are being deciphered, less is known about the precise tissue localization and the way by which each symbiotic partner is transmitted from parent to offspring. The aim of this work was to understand how tsetse endosymbiotic bacteria are transmitted from one generation to the other, by analyzing the precise localization of the endosymbionts in host tissues associated with reproduction. To this end, we have used Fluorescent In Situ Hybridization (FISH) by using specific fluorescent oligoprobes designed to match specifically with 16S rRNA of the different bacteria.

Main results

1. The bacteriome tissue houses Wigglesworthia only As an obligate endosymbiont, Wigglesworthia is contained in bacteria-bearing host cells called the bacteriocytes, which are grouped to form the bacteriome organ. This tissue is located in the anterior midgut forming two lobes surrounding the gut. FISH experiments with the Wigglesworthia probe showed strong signals from bacteriome sections from the female gut. Each bacteriome lobe is formed by different bacteriocyte layers (Balmand et al., 2012 JIP). Remarkably, applying probes of S-symbionts failed to show any signal on the same sections, suggesting a monoinfection status of tsetse bacteriocytes with the primary endosymbiont (P-symbiont) Wigglesworthia.

2. Wigglesworthia does not infect reproductive tissues FISH analysis of the female reproductive tract with Wigglesworthia probe failed to reveal any signal from ovaries and uterus (Balmand et al., 2012 JIP). Surrounding adipocytes were also free of Wigglesworthia . Accordingly, the examination of 194 oocytes, trophocytes and embryos did not show any Wigglesworthia inside these germ cells. However, Wigglesworthia was abundant in the lumen of the mother’s milk gland tubules, while some bacterial cells were also observed in the canal leading to the secretory cell vacuoles, where the milk products are produced and secreted to feed the developing progeny in the uterus. Wigglesworthia could also be found extracellularly in cavities within the fat body in larva. As for the progeny, infection with Wigglesworthia seems to occur late during development. Wigglesworthia was detected in the first instar larvae, at low amounts surrounding the intestine. The bacterial density increased in the bacteriome of the third instar larvae. Interestingly, Wigglesworthia does not infect the larval intestine but remains confined to the second layer of cells that surround the gut making up the bacteriome, similar to that described in Blochmannia association with the carpenter ant Camponotus floridanus . Taken together, these data indicate that Wigglesworthia constitutes two separate sub-populations in adult females: the first strictly intracellular within the bacteriocytes in the adult gut and the second free-living in the lumen of milk glands and inside fat body cavities in larva.

3. Sodalis endosymbionts are in the reproductive tract and infect milk gland cells and lumen. As was the case for Wigglesworthia, no Sodalis cells were detected in oocytes and embryos (Balmand et al., 2012). However, unlike Wigglesworthia , Sodalis cells are spread throughout the reproductive tract. They were shown to infect the oviduct, all over the uterine muscle tissue, in the fat body, and even in spermatheacae sheath cells. Moreover, Sodalis can be detected in the lumen of the milk gland , intracellular within the secretory cells, or in both compartments. In the milk gland lumen, cohabitation between Sodalis and Wigglesworthia was shown is only few cases, indicating a possible competition between these bacteria. In young larval progeny, the S-symbiont Sodalis can be detected in the first instar larvae, as the P-symbiont Wigglesworthia . However, while Wigglesworthia invades the bacteriocyte cells and differentiates into the bacteriome organ, Sodalis cells form aggregate islands throughout the fat body and gut.

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4. Wolbachia infects early oocytes and embryos Wolbachia endosymbiont is the only bacterium that infects embryos prior to intrauterine development. Wolbachia was seen in the developing oocytes, being at both poles as a thin layer, within the yolk, just above the follicular cells. In intimate connection with the oocytes, four out of five trophocytes analyzed were highly infected with Wolbachia . Apart from these cells, Wolbachia was not detected in the fat body and in the uterus. Trophocytes could represent a Wolbachia reservoir, intimately connected to oocytes. When oocytes move down through the uterus to hatch into an embryo, Wolbachia can still be detected at both poles of the embryo, constituting aggregates on the follicular cell layer. Later in second instar larvae, Wolbachia was seen sporadically in aggregates. Contrary to the nutritional endosymbionts Wigglesworthia and Sodalis , Wolbachia was not detected in the milk gland. It is unlikely that milk secretions provide a route of transmission for Wolbachia .

Reference: Severine Balmand, Claudia Lohs, Serap Aksoy, Abdelaziz Heddi (2012 ). Tissue distribution and transmission routes for the Tsetse fly endosymbionts. Journal of Invertebrate Pathology (in press)

University of Western Greece: Kostas Bourtzis Collaborators: Serap Aksoy, Adly Abd-Alla, Peter Takac, Imna Malele, Alexander P. Egyir Yawson, Florence Wamwiri, Anna Malacrida

Abstract Wolbachia is a widespread and highly diverse group of bacterial symbionts infecting arthropod and filarial nematode species. These alphaproteobacteria are frequently found as intracellular and maternally inherited symbionts in insect species, whose reproductive properties they manipulate by inducing cytoplasmic incompatibility (CI), parthenogenesis, male-killing and feminization, thus spreading efficiently and rapidly into host populations. Due to their unique biology, Wolbachia symbiotic associations are currently being considered as promising tools for the control of agricultural pests and medical disease vectors, including Glossina sp., which transmit African trypanosomes, the causative agents of the sleeping sickness in humans and of nagana in animals. In the present study, we report on recent studies on the detection and characterization of Wolbachia infections in Glossina species, the horizontal transfer of Wolbachia genes to tsetse chromosomes and also on how Wolbachia symbiosis could be harnessed for the development of novel symbiont- based pest and disease control strategies.

Introduction Wolbachia is an obligatory intracellular and maternally transmitted endosymbiont which has been associated with the induction of a variety of reproductive abnormalities such as cytoplasmic incompatibility (CI), male-killing, feminization, and parthenogenesis. Wolbachia symbiosis and Wolbachia -induced reproductive alterations have been reported in numerous arthropods hosts, they are, however, most common in insects. Tsetse flies (Glossina spp.) are the sole vectors of Trypanosoma spp., the causative agents of sleeping sickness in humans (human African trypanosomosis, HAT) and of nagana (African animal trypanosomosis, AAT) in livestock. Although epidemics have significantly declined during the last years, still tens of millions of people in Africa continue to be at risk of contracting sleeping sickness. There are several accepted environment-friendly methods to control the insect vector, including the sterile insect technique (SIT). Recently, there is an increasing interest to develop and use Wolbachia -based environment- friendly strategies for the control of insect pests and disease vectors, including tsetse flies and trypanosome transmission. The main goals of our research activities in the frame of this CRP were: (a) the detection and characterization of Wolbachia infections in both laboratory and natural populations of tsetse flies and (b) investigate the possible correlation between Wolbachia and salivary gland hypertrophy virus infections. In this report, we also discuss how Wolbachia Page 31 symbiosis could be harnessed for the control of tsetse flies and trypanosomosis.

Main Results 1. Detection of Wolbachia in laboratory and natural populations of tsetse flies We applied a specific 16S rRNA PCR assay to detect the presence of Wolbachia in a total of 5339 specimens of eleven different Glossina species (G. m. morsitans, G. m. centralis, G. m. submorsitans, G. austeni, G. brevipalpis, G. pallidipes, G. p. palpalis, G. p. gambiensis, G. fuscipes fuscipes, G. tachinoides and G. medicorum ). These specimens consisted of eight laboratory stocks and thirteen natural populations originating from 13 African countries.

2. Characterization of Wolbachia strains from tsetse flies The characterization of tsetse Wolbachia strains was based on Multi Locus Typing System (MLST) and the wsp gene (outer surface protein). We selected a total of 23 Wolbachia -infected Glossina specimens from different areas and species to be genotyped by amplifying the five MLST housekeeping genes ( gatB, coxA, hcpA, ftsZ and fbpA ), as well as the wsp gene. So far, 12 allelic profiles or Sequence Types (ST) were detected in tsetse flies Wolbachia strains. Moreover, the same 23 samples were genotyped using the wsp (Wolbachia surface protein) gene, for which 15 alleles were found. We also identified their WSP HVR profile (a combination of the four HVR amino acid haplotypes). To date, a total of fourteen WSP HVR profiles were found, of which twelve were new entries to the Wolbachia WSP database. We also constructed four phylogenetic trees based on: a) the concatenated data set of the MLST Wolbachia genes, b) the Wolbachia wsp gene, c) the Wolbachia 16S rRNA gene and d) the mitochondrial cytochrome oxidase subunit I which clearly suggest that Glossina species have experienced multiple Wolbachia invasions during evolution.

3. Horizontal transfer of Wolbachia genes to tsetse chromosomes During the 16S rRNA screening and the MLST analysis, evidence for horizontal transfer of the 16S rRNA, gatB, coxA, hcpA, ftsZ, fbpA and wsp genes to the host chromosomes was observed in specimens of three Glossina species ( G. morsitans morsitans, G. pallidipes and G. austeni ). Cloning and sequencing analysis showed that the horizontally transferred Wolbachia genes were pseudogenized.

4. Wolbachia genomics – Wolbachia ankyrins We developed in silico strategies to “fish out” Wolbachia-specific sequences through the whole genome sequence project of G. m. morsitans. Two main horizontal genomic transfer events were detected, fully annotated and were compared with the genome of the cytoplasmic Wolbachia strain sequenced by 454 technology at Prof. Alsoy’s laboratory. The genome size of the tsetse Wolbachia chromosomal insertions is reduced and has fewer CDSs, shorter genes, an increased number of pseudogenes for the chromosomal insertions, as expected. The majority of the chromosomal CDSs are pseudogenes, they contain a lot of SNPs, as well as several deletions or insertions, which render these CDSs inactive. Several characteristic features of the Wolbachia genome are retained, such as the presence of ankyrin genes and prophages, ISs etc.

5. Wolbachia and salivary gland hypertrophy virus The specimens were also screened for the presence of the tsetse salivary gland hypertrophy virus (Gp -SGHV). This screen suggests that there is a negative correlation between the presence of Wolbachia and SGHV.

6. Wolbachia -induced CI - Wolbachia -based control strategies Wolbachia -induced CI has been proposed as a potential mechanism for the control of agricultural pests and disease vectors. Our study shows that Wolbachia is present in both laboratory and natural populations of tsetse flies. In addition, Alam and colleagues (2011) recently reported that the presence of Wolbachia can induce strong CI in G. m. morsitans . This finding opens new routes for

Page 32 population control of the insect vector and of trypanosomosis. This can be done with three potential approaches: (a) to use Wolbachia -induced CI as a population suppression mechanism in a way analogous to SIT; (b) to use Wolbachia -induced CI as a spreading/replacement mechanism for desired phenotypes and (c) Wolbachia -induced CI can also be used to drive trypanosome resistant paratransgenic tsetse into natural populations to replace their parasite-susceptible counterparts.

Publications relevant to CRP P. Ioannidis and K. Bourtzis (2007). Insect Symbionts and Applications: the paradigm of cytoplasmic incompatibility-inducing Wolbachia. Entomological Research , 37: 125-138. N. Lo, C. Paraskevopoulos, K. Bourtzis, S. L. O’Neill, J. H. Werren, S. R. Bordenstein and C. Bandi (2007). Taxonomic status of the intracellular bacterium Wolbachia pipientis . International Journal of Systematic and Evolutionary Microbiology, 57: 654-657. P. Ioannidis, J.C. Dunning Hotopp, P. Sapountzis, S. Siozios, G. Tsiamis, S.R. Bordenstein, L. Baldo, J.H. Werren and K. Bourtzis (2007). New Criteria for Selecting the Origin of DNA Replication of Wolbachia and Closely Related Bacteria. BMC Genomics , 8:182. P. Ioannidis and K. Bourtzis (2007). Insect Symbionts and Applications: the paradigm of cytoplasmic incompatibility-inducing Wolbachia. Entomological Research , 37: 125-138. Bourtzis, K., 2008. Wolbachia -based technologies for insect pest population control. Adv Exp Med Biol. 627 , 104-13.38. S. Siozios, P. Sapountzis, P. Ioannidis and K. Bourtzis (2008). Wolbachia Symbiosis and Insect Immune Response. Insect Science , 15: 89-100. S. Zabalou, A. Apostolaki, S. Pattas, Z. Veneti, C. Paraskevopoulos, I. Livadaras, G. Markakis, T. Brissac, H. Merçot and K. Bourtzis (2008). Multiple rescue factors within a Wolbachia strain. Genetics , 178: 2145-2160. S.R. Bordenstein, C. Paraskevopoulos, J.C. Dunning-Hotopp, P. Sapountzis, N. Lo, C. Bandi, H. Tettelin, J.H. Werren and K. Bourtzis (2009). Parasitism and mutualism in Wolbachia : what the phylogenomic trees can and can not say. Molecular Biology and Evolution , 26: 231-241. L. Klasson, J. Westberg, P. Sapountzis, K. Näslund, Y. Lutnaes, A. C. Darby, Z. Veneti, L. Chen, H. R. Braig, R. Garrett, K. Bourtzis and S. G. E. Andersson (2009). The mosaic genome structure of the Wolbachia wRi strain infecting Drosophila simulans. Proceedings of the National Academy of Sciences of the United States of America, 106: 5725-5730. N. Ishmael, J.C. Dunning-Hotopp, P. Ioannidis, S. Biber, J. Sakamoto, S. Siozios, V. Nene, J. Werren, K. Bourtzis, S.R. Bordenstein, H. Tettelin (2009). Extensive Genomic Diversity of Closely Related Wolbachia Strains. Microbiology SGM , 155: 2211-2222. S. Zabalou, A. Apostolaki, I. Livadaras, G. Franz, A.S. Robinson, C. Savakis and K. Bourtzis (2009). Incompatible Insect Technique: Incompatible Males from a Ceratitis capitata (Diptera: Tephritidae) Genetic Sexing Strain. Entomologia Experimentalis et Applicata , 132: 232-240. R. Gross, F. Vavre, A. Heddi, G.D. Hurst, E. Zchori-Fein and K. Bourtzis (2009). Immunity and Symbiosis. Molecular Microbiology , 73: 751-759. A. Saridaki and K. Bourtzis (2009). Wolbachia -induced reproductive parasitism and applications. Entomologia Hellenica , 18: 3-16. A. Saridaki and K. Bourtzis (2010). Wolbachia : more than just a bug in insects’ genitals. Current Opinion in Microbiology 13: 67-72. Luciano Sacchi, Marco Genchi, Emanuela Clementi, Ilaria Negri, Alberto Alma, Davide Sassera, Stefan Oehler, Kostas Bourtzis and Claudio Bandi (2010).Bacteriocyte-like cells harbour Wolbachia in the ovary of Drosophila melanogaster (Insecta, Diptera) and Zyginidia pullula (Insecta, Hemiptera). Tissue & Cell 42: 328-333. A. Apostolaki, A. Saridaki, I. Livadaras, C. Savakis and K. Bourtzis (2011). Transinfection of the olive fruit fly with a Wolbachia CI inducing strain: a promising symbiont-based population control strategy? Journal of Applied Entomology 135: 546–553 (doi: 10.1111/j.1439- 0418.2011.01614.x). A. Saridaki, P. Sapountzis, H. L. Harris, P. D. Batista, J. A. Biliske, H. Pavlikaki, S. Oehler, C. Savakis, H. R. Braig and K. Bourtzis (2011). Wolbachia Prophage DNA Adenine Page 33

Methyltransferase Genes in Different Drosophila-Wolbachia Associations. PLoS ONE 6(5) : e19708 (doi:10.1371/journal.pone.0019708). G. Papafotiou, S. Oehler, C. Savakis and K. Bourtzis (2011). Regulation of Wolbachia ankyrin- domain encoding genes in Drosophila gonads. Research in Microbiology 162:764–772 (doi: 10.1016/j.resmic.2011.06.012). V. Doudoumis, G. Tsiamis, F. Wamwiri, C. Brelsfoard, U. Alam, E. Aksoy, S. Dalaperas, A. Abd- Alla, J. Ouma, P. Takac, S. Aksoy and K. Bourtzis (2012). Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse (genus Glossina) . BMC Microbiology , 12 (Suppl 1): S3. V. Doudoumis, U. Alam, E. Aksoy, A. Abd-Alla, G. Tsiamis, C. Brelsfoard, S. Aksoy and K. Bourtzis (2012). Tsetse-Wolbachia Symbiosis: comes of age and has great potential for pest and disease control. Journal of Invertebrate Pathology (In Press). A. Abd-Alla, M. Bergoin, A. G. Parker, N. K. Maniana, J. M. Vlak, K. Bourtzis, D. G. Broucias and S. Aksoy (2012). Improving SIT for tsetse flies through research on their symbionts and pathogens. Journal of Invertebrate Pathology (In Press). J. Van den Abbeele, K. Bourtzis, B. Weiss, C. Cordón-Rosales, W. Miller, A. Abd-Alla and A. G. Parker, (2012). Enhancing Tsetse fly refractoriness to Trypanosome infection - A new IAEA Coordinated Research Project. Journal of Invertebrate Pathology (In Press).

Institute of Tropical Medicine Antwerp, Belgium: Van Den Abbeele, J*. Collaborators: Aksoy, S.

Abstract

During this CRP we have developed a suitable expression system for heterologous proteins in cultured Sodalis . We demonstrated that the twin-arginine (TAT) translocation pathway in Sodalis is active to export active heterologous proteins to the periplasm. In addition, we showed that the pelB leader peptide is suitable to direct the export of functional anti-trypanosome single domain antibody (Nanobody ®) to the periplasm of S. glossinidius resulting in significant levels of extracellular nanobody release. These S odalis strains are shown not to be affected in their growth compared to the wildtype Sodalis , suggesting that they may be competitive with endogenous microbiota in the midgut environment of the tsetse fly. These data are the first demonstration of the expression and extracellular release of functional trypanosome-interfering proteins in S. glossinidius , further supporting its use as a paratransgenic platform organism

Introduction

Sodalis glossinidius has a wide host tissue tropism, and is found intra- and extracellularly in tsetse’s gut, haemolymph and salivary glands. The function of this bacterium in tsetse is speculative, although previous research suggests it plays a role in host longevity and susceptibility to trypanosome infection. This recently acquired symbiont is an ideal candidate for use in a disease control strategy called ‘paratransgenesis’. This strategy involves culturing Sodalis outside of tsetse and then genetically modifying the cells to produce anti-trypanosome proteins/peptides. These genetically modified Sodalis bacteria can then be re-introduced to re-populate the fly. The gut of these flies is then colonized with rec Sodalis , where they subsequently cohabit with pathogenic trypanosomes. The anti-trypanosome proteins delivered by the rec Sodalis will interfere with parasite development in the tsetse midgut. In the context of this paratransgenesis approach we aimed to develop in this CRP an expression system for Sodalis that allows for the expression and secretion of anti-trypanosome molecules

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Main Results .

1. Functional analysis of the Twin-Arginine Translocation Pathway in Sodalis glossinidius , a bacterial symbiont of the tsetse fly This study demonstrates a functional twin-arginine (TAT) translocation pathway present in the tsetse fly symbiont Sodalis glossinidius and its potential to export active heterologous proteins to the periplasm. So far, the functionality of the bacterial Tat system has been analyzed mainly in free- living organisms. We showed that the Tat pathway is biologically active in Sodalis glossinidius , a bacterial symbiont of the tsetse fly, and identified a number of predicted natural endogenous substrates that contain the characteristic twin-arginine motif. Functionality was demonstrated using green fluorescent protein (GFP) fused to the Tat signal peptide of E. coli trimethylamine N-oxide reductase (TorA). Moreover, we illustrated that the Tat system has the potential to be exploited for exporting heterologous proteins in an active manner to the periplasm.

2. Expression and extracellular release of a functional anti-trypanosome Nanobody ® in Sodalis glossinidius , a bacterial symbiont of the tsetse fly In this study, S. glossinidius was transformed to express a single domain antibody, (Nanobody ®) Nb_An33, that efficiently targets conserved cryptic epitopes of the variant surface glycoprotein (VSG) of the parasite Trypanosoma brucei . Next, we analyzed the capability of two predicted secretion signals to direct the extracellular delivery of significant levels of active Nb_An33. We showed that the pelB leader peptide was successful in directing the export of fully functional Nb_An33 to the periplasm of S. glossinidius resulting in significant levels of extracellular release. Finally, S. glossinidius expressing pelBNb_An33 exhibited no significant reduction in terms of fitness, determined by in vitro growth kinetics, compared to the wild-type strain. These data are the first demonstration of the expression and extracellular release of functional trypanosome-interfering Nanobodies ® in S. glossinidius . Furthermore, Sodalis strains that efficiently released the effector protein were not affected in their growth, suggesting that they may be competitive with endogenous microbiota in the midgut environment of the tsetse fly. Collectively, these data reinforce the notion for the potential of S. glossinidius to be developed into a paratransgenic platform organism.

Summary

• Development of an expression system for heterologous proteins in cultured Sodalis . • Demonstration that the twin-arginine (TAT) translocation pathway in Sodalis is active to export active heterologous proteins to the periplasm. • Demonstration that the pelB leader peptide is suitable to directthe export of functional anti- trypanosome single domain antibody (Nanobody ®) to the periplasm of S. glossinidius resulting in significant levels of extracellular nanobody release. • These S odalis strains are shown not to be affected in their growth compared to the wildtype Sodalis , suggesting that they may be competitive with endogenous microbiota in the midgut environment of the tsetse fly.

Publications

De Vooght L., Caljon G., Coosemans M., Van Den Abbeele J. (2011) Functional analysis of the Twin-Arginine Translocation Pathway in Sodalis glossinidius , a bacterial symbiont of the tsetse fly. Applied and Environmental Microbiology 2011, 77(3):1132-4

De Vooght L., Caljon G., Stijlemans B., De Baetselier P., Coosemans M., Van Den Abbeele J. (2012) Expression and extracellular release of a functional anti-trypanosome Nanobody ® in Sodalis glossinidius , a bacterial symbiont of the tsetse fly. Microbial Cell Factories 2012, 11:23 Page 35

Caljon G., De Vooght L.,Van Den Abbeele J. (-) Options for the delivery of anti-pathogen molecules in arthropod vectors. Journal of Invertebrate Pathology , special issue.

OVI/ARC, Onderstepoort, South Africa: Koekemoer, O. Collaborators: Abd Alla, A., Vlak, J., Aksoy, S., Malacrida, A.

Abstract

Field samples of G. brevipalpis and G. austeni were collected from areas covering the total known distribution of the two species in South Africa and from the Mlawula National park in Swaziland. DNA extractions were made from whole flies and used to determine the presence of both Wolbachia and GpSGHV by PCR. The GpSGHV-specific PCR revealed only a single infection in colonized flies and very low levels of infection in wild specimens. G. austeni had an average Wolbachia infection rate of 95% while G. brevipalpis only showed infection in 61%. Typing of the symbiont from Wolbachia positive flies using the MLST approach revealed the presence of at least two genotypes in G. brevipalpis , one of which has a new sequence on the hcpA locus. Further genetic distinctions could be made between Wolbachia infecting G. brevipalpis and G. austeni from different locations using a high resolution melting profile analysis of wsp -gene PCR products.

Summary of Results

The total area in South Africa and one known location in Swaziland as well as colony tsetse were surveyed for the presence and signs of SGHV and Wolbachia in G. austeni and G. brevipalpis using PCR methods and physical examinations.

Except for one G. brevipalpis , there was no positive result when testing tsetse from the colonies at Onderstepoort. One set of degenerate primer (pif-2) revealed results that might indicate the presence of viral DNA. This will have to be confirmed using sequencing of the PCR products as p74-based PCR revealed no positives. This might be the result of nucleotide variations in the sites where the PCR-primers are to bind. No physical evidence of salivary gland hypertrophy could be observed in any case where freshly collected flies were examined.

The Wolbachia surveys revealed a very high rate of infection in G. austeni . In multiple areas the infection rate was 100%. The lowest rate of infection in this species was 83%. In G. brevipalpis there was much bigger variation ranging from areas that showed a 0% infection rate to areas where 91% of the flies were positive. Selected positive individuals were used to carry out molecular typing of their Wolbachia symbionts using two different assays. In the first analysis the MLST protocol of Baldo et al., (2006) was used. Results from G. austeni were the same as recently reported by Doudoumis et al. (2012). No variation was observed between different origins, no new alleles could be detected except for one seemingly new allele for the CoxA locus of a colony fly. G. brevipalpis , however, showed variation in one set of samples from the southern part of the Hluhluwe Imfolozi Park. Sequence variation was observed on all 5 loci between Wolbachia alleles from these flies and specimens from other locations in South Africa. Furthermore, flies from this location showed a new allele, not currently present in the MLST database on the hcpA gene.

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A paper by Henri and Mouton (2011) describes a methods that makes use of PCR and HRM to rapidly analyse Wolbachia for genetic diversity. It does not rely on sequencing and targets the highly variable wsp-gene. One of the regions on this gene, HVR3, showed genetic diversity within the Wolbachia populations that were tested, and revealed variation between flies that were not seen using the MLST approach. In the case of G. brevipalpis a distinction could be seen between two groups of flies, one containing specimens from the Kosi bay, Hellsgate and Hluhluwe areas, and the other specimens from Tembe, Hellgate, False Bay park and Charter’s Creek. This not only confirms the genetic diversity that was observed with the MLST analysis, but it shows that there is variation within at least one area viz. Hellsgate. Wolbachia from G. austeni from Swaziland showed no genetic variation using MLST but a clear variation was observed in one specimen from this location. Again, there was variation within the group, as other specimens from Swaziland showed the same profile as that of the Wolbachia that were from flies collected in South Africa.

In summary, although the virus-specific PCR showed some results that can be interpreted as positive for the presence of the virus, this needs to be confirmed by sequencing of the amplicons. Wolbachia infection rates were established for the different areas under survey and genetic typing has revealed genetic variation in symbionts from both host species occurring in South Africa and Swaziland.

Institute of Zoology, Slovak Academy of Sciences. Slovakia: Takac, P. Collaborators: Aksoy, S., Malacrida, A., Abd-Alla, A.

Abstract During the CRP project and within the contract agreement we focused on the provision of tsetse biological material to research groups. We had been supply of 10,000 tsetse fly puparia as required to participants of the CRP D42012 "Improving SIT for tsetse flies through research on their symbionts and pathogens in 14 shipments per year. a. 200 puparia shipped from G. m. morsitans, G. pallidipes and G. f. fuscipes colonies monthly to S. Aksoy, Yale. b. Four allotments of 200 puparia from G. m. morsitans shipped to A. Malacrida, Pavia. c. Puparia (1000 for each species) from G. pallidipes and G. fuscipes shipped to N. Maniana, ICIPE. At the same time we set up the collaborative experiments with Aksoy´s lab., concerning the supplementation of the host diet with yeast , different vitamine mixtures and different antibiotics, to study the influence of this treatment to microbial flora, fecundity and mortality of G. morsitans morsitans and G. fuscipes fuscipes colonies.

Introduction Glossinidae (Diptera) are vectors of African pathogenic trypanosomes, which are of medical and economic importance. They are exclusively haematophagous and this higly restricted nutritional ecology has resulted in obligate adaptations with symbiotic bacteria in tsetse. The endosymbionts provide nutritional supplements, in the absence of which females are becoming sterile. Tsetse´s dependence on its obligate microbiota for reproduction provides a weak link in its biology, and may generate alternative control strategies that should be explored. These endosymbionts have also an important role in parasite transmission. The mutual interactions result in host immune regulation and nutritional environment in the midgut and may also contribute to the vector susceptibility traits observed in field populations. This way, the endosymbionts may be potentially an efficient target for controlling tsetse fly vectorial competence and consequently sleeping sickness. The three organisms have been already characterized for Glossina genus, Wigglesworthia glossinidia, Sodalis glossinidius and Wolbachia pipientis and all three symbionts are, in essence, materially associated with the metabolism of B-complex vitamins essential for tsetse survival, the effect of yeast based meals and different B-complex vitamins meals on fitness and fecundity was analyzed.

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Results 1. Provision of biological tsetse material to CRP research groups During the CRP project and within the contract agreement we focused on the provision of tsetse biological material to research groups. We had been supply of 10,000 tsetse fly puparia as required to participants of the CRP D42012 "Improving SIT for tsetse flies through research on their symbionts and pathogens in 14 shipments per year. a. 200 puparia shipped from G. m. morsitans, G. pallidipes and G. f. fuscipes colonies monthly to S. Aksoy, Yale. b. Four allotments of 200 puparia from G. m. morsitans shipped to A. Malacrida, Pavia. c. Puparia (1000 for each species) from G. pallidipes and G. fuscipes shipped to N. Maniana, ICIPE.

2. Impact of antibiotic, yeast and yeast vitamims mixture treatment on host fecundity and lifespan We evaluated the effect of continuous per os treatment of fertile females with the tetracycline and ampiciline based antibiotics and yeast extracts or vitamins mixture diets by measuring the total number of larvae deposited per group over the course of three deposition cycles. To determine if the tetracycline and ampiciline antibiotics have a prolonged effects on the process of metamorphosis within the larvae or pupae. The hatching rate of progeny was determined in these groups. At the same time we measured the influence of antibiotic and yeast treatment on fly lifespan. We measured the effect of tetracycline, tetracycline and yeast, ampiciline, ampiciline and yeast ; only yeast and different vitamine mixture treatment on pupae production. Under optimum conditions the first gonotrophic cycle takes about 20–22 days for development from egg to parturition. In subsequent gonotrophic cycles females produce a larva every 9 to 11 days. Ampiciline treatment does not reduce fecundity since it does not damage Wigglesworthia resident within bacteriocytes in the midgut, unlike tetracycline, which clears all bacteria including Wigglesworthia and Wolbachia and induces sterility. Accordingly, ampicillin-receiving flies remained fecund while tetracycline receiving flies were rendered sterile. Yeast extract (10% w/v) provisioning of the blood meal rescued fecundity of the females receiving tetracycline to similar levels as that of wild type and ampicillin receiving flies (65%, 55% and 64% over the first gonotrophic cycle and 53%, 58% and 49% over the second gonotrophic cycles, respectively). However, yeast provisioning at 10% w/v had a cost on fecundity when compared to flies maintained on normal blood meals, (92% versus 55% over the first gonotrophic cycle and 92% and 58% over the second gonotrophic cycle, respectively). Nevertheless, yeast supplementation was able to rescue the tetracycline-induced sterility to levels comparable to those observed for Gmm Wt receiving yeast or ampicillin supplemented blood meals, respectively.Thus yeast supplemented dietary regiment allowed us to develop two lines to analyze the functional role of Wolbachia symbionts in tsetse biology; one lacking all symbionts ( Gmm Apo ) and another lacking Wigglesworthia but still retaining Wolbachia and Sodalis (Gmm Wig−). The Gmm Apo progeny resulting from the first and second depositions of tetracycline treated mothers were tested for the presence of Sodalis, Wigglesworthia and Wolbachia by a bacterium-specific PCR-assay. The PCR-assay demonstrated the absence of all three symbionts as early as the first deposition in both the male and female Gmm Apo adults. The absence of Wolbachia from the reproductive tissues of Gmm Apo females was also verified by Fluorescent In Situ Hybridization (FISH) analysis. In contrast, Wolbachia was present in egg chambers during both early and late developmental stages in Gmm Wt females. Longevity of F1 Gmm Apo females was compared to that of Gmm Wt adults maintained on the same yeast-supplemented blood meal over 40 days (two-gonotrophic cycles). No difference (X 2 = 0.71, df = 1, P = 0.4) was observed in survivorship comparisons between the two groups. The second line (Gmm Wig−) generated from ampicillin treated females still retain their Wolbachia and Sodalis symbionts, while lacking both Wigglesworthia populations as evidenced by FISH and PCR amplification analysis. When maintained on yeast-supplemented blood, this line (similar to

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Gmm Apo ) also did not display any longevity differences from the Gmm Wt adults sustained on the same diet.

References Alam U , Medlock J , Brelsfoard C , Pais R , Lohs C , Balmand S , Carnogursky J , Heddi A , Takac P , Galvani A , Aksoy S. (2011) Wolbachia Symbiont Infections Induce Strong Cytoplasmic Incompatibility in the Tsetse Fly Glossina morsitans. PLoS Pathog 7(12): e1002415. doi:10.1371/journal.ppat.1002415. http://www.plospathogens.org/article/info%3Adoi%2F10.1371%2Fjournal.ppat.1002415;jsessio nid=554224F4111AA0B7AB53D29EC2D2BB5A

DOUDOUMIS V, TSIAMIS G, WAMWIRI F, BRELSFOARD C, ALAM U, AKSOY E, DALAPERAS S, ABD-ALLA Adly, OUMA J, TAKAC P, AKSOY S, BOURTZIS K. (2012) Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina). BMC Microbiology, vol. 12, suppl. 1, s3; 13pp. ISSN 1471-2180. http://www.biomedcentral.com/1471-2180/12/S1/S3

Austria, Wolfgang J. Miller Collaborators: Adly M. M. Abd-Alla, Andrew G. Parker

Abstract In the course of this CRP we demonstrate the high applicability of a novel multi-copy IS-based (Insertion Sequences) and VNTR-based (Variable-Number-Tandem-Repeat) molecular screening tool for fingerprinting Wolbachia -infections in tsetse flies. The VNTR-141 locus provides reliable and concise differentiation between Wolbachia strains deriving from Glossina morsitans morsitans, Glossina morsitans centralis , and Glossina brevipalpis. Moreover, we show that certain Wolbachia- infections in Glossina spp. are capable of escaping standard PCR screening methods by ‘hiding’ as low-titer infections below the detection threshold. By applying a highly sensitive PCR-blot technique to our Glossina specimen, we were able to enhance the symbiont detection limit substantially and, consequently, trace unequivocally Wolbachia -infections at high prevalence in laboratory-reared G. swynnertoni individuals. To our knowledge, Wolbachia -persistence was reported exclusively for field-collected samples, and at low prevalence only. Finally, we demonstrated the substantially higher Wolbachia titer levels found in hybrid Glossina compared to non-hybrid hosts and the possible impact of these titers on hybrid host fitness that potentially trigger incipient speciation in tsetse flies.

Results

By the optimization and application of our improved Wolbachia detection tools, i.e., (i) IS transposons, (ii) VNTR-fingerprinting, (iii) wsp blot-PCR, and (iv) artificial titer enhancement in hybrids we have succeeded in uncovering low-titer infections in tsetse fly species and lab colonies that were earlier overlooked by standard Wolbachia detection methods. Furthermore, by means of our hypersensitive detection tools we have assigned much higher infection frequencies than earlier reported in tsetse flies from field populations and lab colonies. In addition to the power of this high-resolution method we have also demonstrated the capacity of tsetse fly Wolbachia to induce CI in hybrids between Glossina species that presumably triggers incipient speciation in tsetse flies. According to previous studies, barriers to hybrid formation in the genus Glossina are weak since natural hybridization between the species has been reported repeatedly (reviewed in Gooding 1990). In this study, however, we have observed barriers to Page 39 hybridization on both post- and pre-mating levels. Interestingly, Gooding has suggested in one of his recent publications (Gooding and Krafsur, 2005) that mating barriers are already in the process of formation and that tsetse flies could serve as a suitable system for studying ongoing speciation. As in the situation observed in hybrids between the semispecies of the D. paulistorum species cluster (Miller et al. 2010), Glossina hybrids are prone to male sterility and reduced fecundity of females (Gooding 1993). Here, we show that Wolbachia titers are massively increased in most F1 hybrids and speculate that this over-replication could trigger the aforementioned strong post-mating isolation. We suggest that we probably uncovered the induction of Wolbachia -induced CI in cross breeding of morsitans group taxa, causing high levels of early embryonic and/or late pupal lethality. Regarding the finding that both parents harbor Wolbachia at high prevalence and that F1 hybrid formation is affected in both directions, not only uni- but also bidirectional CI in tsetse flies is quite likely. As earlier demonstrated in Culex pipiens , parasitoid wasps of the genus Nasonia and neotropical Drosophila species expression of strong bidirectional CI can be regarded as effective post-mating barriers capable of fostering incipient host speciation (Yen and Barr 1971; Bordenstein et al. 2001; Miller et al. 2010). The manuscript reporting our new data is accepted for publication as an original paper in the Journal of Invertebrate Pathology as Schneider DI, Garschall KI, Parker AG, Abd-Alla AMM, Miller WJ (2012) entitled Global Wolbachia prevalence, titer fluctuations and their potential of causing cytoplasmic incompatibilities in tsetse flies and hybrids of Glossina morsitans subgroup species.

Summary

• Successful application and optimization of candidate IS and VNTRs primer sets on diverse Glossina samples from lab colonies and field populations. • Clone & sequence informative candidate loci of Wolbachia marker genes from different tsetse species • Establish hyper-sensitive wsp-Blot PCR for detecting low-copy Wolbachia in tsetse • Determine Wolbachia low titer infections & multiple infections in variant tsetse flies • Monitoring Wolbachia dynamics in tsetse hybrids (knock-in)

Publication : Schneider DI, Garschall KI, Parker AG, Abd-Alla AMM, Miller WJ (2012) entitled Global Wolbachia prevalence, titer fluctuations and their potential of causing cytoplasmic incompatibilities in tsetse flies and hybrids of Glossina morsitans subgroup species. JIP in press.

References:

Bordenstein S.R., et al. (2001) Wolbachia -induced incompatibility precedes other hybrid incompatibilities in Nasonia . Nature 409, 707-710. Gooding, R.H., 1990. Postmating barriers to gene flow among species and subspecies of tsetse flies (Diptera: Glossinidae). Can. J. Zool. 68, 1727-1734. Gooding, R.H., 1993. Hybridization of Glossina swynnertoni with subspecies of Glossina morsitans (Diptera: Glossinidae): implications for use of hybrid sterility and satyrs for genetic control of tsetse, in: Proceedings of the International Symposium on Management of Insect Pests: Nuclear and Related Molecular and Genetic Techniques, Vienna, 19-23 October 1992. IAEA, Vienna, pp. 603-617. Miller, W.J., et al., 2010. Infectious speciation revisited: impact of symbiont-depletion on female fitness and mating behavior of Drosophila paulistorum . PLoS Pathog. 6, e:1001214. Riegler. M., et al., 2012. Tandem repeat markers as novel diagnostic tools for high resolution fingerprinting of Wolbachia . BMC Microbiol. 12(Suppl 1):S12

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Schneider DI, Garschall KI, Parker AG, Abd-Alla AMM, Miller WJ (2012) entitled Global Wolbachia prevalence, titer fluctuations and their potential of causing cytoplasmic incompatibilities in tsetse flies and hybrids of Glossina morsitans subgroup species. JIP in press. Yen, J.H., Barr, A.R., 1971. New hypothesis of the cause of cytoplasmic incompatibility in Culex pipiens L. Nature. 232, 657-658.

University of Yaounde I, BP 825, Yaounde , Cameroon: Flobert Njiokou Collaborators: Anne Geiger, Adly abd alla Abstract: In order to study the relation between the presence of the symbiont Sodalis glossinidius and the vectorial competence of tsetse flies, entomological and parasitological investigations were carried on in three HAT foci (Bipindi 2008 and 2011, Campo 2008 and Fontem 2009) in Cameroon. Tsetse flies were trapped at two seasons in three villages of each HAD focus. Tsetse fly samples were dissected, the midguts were checked with a microscope for the presence of live trypanosomes, and were subsequently kept in 95% ethanol for the identification of the presence of the symbiont and the trypanosomes using PCR methods. The statistical test for association were use to study the relation between symbiont and trypanosomes. Only Glossina palpalis palpalis was caught in Bipindi and Fontem while three other species were found in Campo dominated by G. p. palpalis (94%). The density of the flies and their infection rate varied among the foci, the villages and the season. The prevalences of Sodalis glossinidus varied among the foci and the villages and were generally high (45 to 69%) whereas trypanosome prevalences significantly differed among the foci, with very low values in Bipindi in 2011. The statistical test revealed significant associations between the symbiont Sodalis glossinidius and many parasite species, in particular, T. b. gambiense T. congolense forest type and the mixed trypanosome infections in the 2008 and 2009 samples. The lack of significant association between Sodalis and the Bipindi 2011 sample of tsetse flies may be due to the very low prevalence of the trypanosome species found due to the HAT control activities.

Introduction The bacterial Sodalis glossinidius was considered to be involved in Glossina vectorial competence to trypanosomes by favouring their installation in the fly midguts. The 14320 Contract was aimed to verify the correlation between the presence of this symbiont in wild tsetse flies and their ability to be naturally infected by trypanosomes.

Main results: -Glossina palpalis palpalis was the more prevalent (94-100% frquency) tsetse fly species in the three HAT foci sampled. -Tsetse flies in the three foci harboured the symbiont Sodalis glossinidius with a rate range between 45 to 60%. Tsetse flies rate of infection by human and animal trypanosomes were relatively high except in the last sample from Bipindi 2011. -Significant statistical correlations were found between S. glossinidius and trypanosome infections. Page 41

Publications: Farikou O., Njiokou F ., Mbida Mbida J.A., Njitchouang G.R., Nana Djeunga H.C., Tazoacha A., Simarro P.P., Cuny G. & Geiger A. 2010. Tripartite interactions between tsetse flies, Sodalis glossinidus and trypanosomes – An epidemiological approach in two historical human African trypanosomiasis foci in Cameroon. Infection, Genetics and Evolution . 10: 115-121.

Farikou O., Njiokou F ., Simo G., Tazoacha A., Cuny G. & Geiger A. Tsetse fly blood meal modification and trypanosome identification in two sleeping sickness foci in the forest of southern Cameroon. 2010. Acta Tropica , 116: 81-88.

UMR 177, IRD-CIRAD, CIRAD TA A-17/G, Campus International de Baillarguet Anne Geiger Collaborators: F. Njiokou

Abstract In a previous work, a large tsetse fly sampling campaign had been conducted in Bipindi and Campo that are two historical sleeping sickness foci located in the south of Cameroon. The results evidenced large differences between the population of flies from Campo and that from Bipindi. So we hypothesized that the geographical isolation of the two foci could have induced independent evolution of fly, and symbiont populations, leading to a divergent diversification of genotypes. To test this hypothesis, we investigated the symbiont genetic structure using the allelic size variation at four specific microsatellite loci. The results showed that the structure of genetic diversity varied at different geographical scales, with almost no differentiation within the Campo Human African Trypanosomiasis (HAT) focus and a low but significant differentiation between the Campo and Bipindi HAT foci. So, the data provided new information on the genetic diversity of S. glossinidius populations. Possible interactions between S. glossinidius subpopulations and Glossina species that could favor tsetse fly infections by a given trypanosome species should be further investigated.

Introduction Bacterial Sodalis glossinidius was considered to be involved in Glossina vectorial competence to trypanosomes by favouring their installation in the fly midguts. The aim of the Research Agreement N° 14266 was to try to decipher the tripartite interaction between tsetse, specific symbiont genotype and trypanosomes in HAT foci of Cameroon.

Main Results - Genetic diversity in S. glossinidius The complete dataset included multilocus genotypes for the 244 S. glossinidius strains from the 244 G. palpalis palpalis sampled in HAT foci in Cameroon (113 from Bipindi, 131 from Campo). The four microsatellite loci used were polymorphic, and a total of 19 alleles were detected, ranging from three (ADNg 12/13) to six (ADNg 21/22, ADNg 15/16) alleles per locus. The combination of the microsatellite alleles yielded a total of 35 haplotypes. The four populations were polymorphic, showing 12-20 haplotypes.

- Population differentiation patterns The population structure of S. glossinidius was explored at different hierarchical levels using AMOVA. No overall differentiation between populations within the Campo focus was observed.

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However, a significant differentiation between the Bipindi (Ebimimbang) population and those of the three villages (Akak, Campo Beach/Ipono, and Mabiogo) in the Campo focus was obtained.

Publications Farikou O., Njiokou F ., Mbida Mbida J.A., Njitchouang G.R., Nana Djeunga H.C., Tazoacha A., Simarro P.P., Cuny G. & Geiger A . 2010. Tripartite interactions between tsetse flies, Sodalis glossinidus and trypanosomes – An epidemiological approach in two historical human African trypanosomiasis foci in Cameroon. Infection, Genetics and Evolution . 10: 115-121. Farikou O., Njiokou F ., Simo G., Asonganyi T., Cuny G. & Geiger A . 2010. Tsetse fly blood meal modification and trypanosome identification in two sleeping sickness foci in the forest of southern Cameroon. Acta Tropica , 116:81-88. Farikou O, Thevenon S, Njiokou F , Allal F, Cuny G, Geiger A . 2011. Genetic Diversity and Population Structure of the Secondary Symbiont of Tsetse Flies, Sodalis glossinidius , in Sleeping Sickness Foci in Cameroon. PLoS Negl Trop Dis .,5(8):e1281. Geiger A , Fardeau ML, Njiokou F , Joseph M, Asonganyi T, Ollivier B, Cuny G. 2011. Bacterial Diversity Associated with Populations of Glossina spp. from Cameroon and Distribution within the Campo Sleeping Sickness Focus. Microb Ecol ., 62(3):632-43. Farikou O, Njiokou F , Cuny G, Geiger A. 2011. Microsatellite genotyping reveals diversity within populations of Sodalis glossinidius, the secondary symbiont of tsetse flies.Vet Microbiol., 150(1- 2):207-10.

University of Pavia, Italy: Malacrida, A *. Collaborator: Aksoy, S., Bourtzis, K., Malele, I., Ouedraogo, G., Koekemoer O., Wamwari, F., Abd-Alla, A.

Abstract We have considered the relations between polyandry, migration rate and the spreading of Wolbachia in wild populations of G. fuscipes and G. pallidipes . Polyandry is a common event in these species and females store sperm from multiple males, potentially available for insemination. Considering that Wolbachia is present in populations of these two species and a certain degree of male migration has been assessed expecially for G. pallidipes, female mate choice may have an impact on the spreading of this endosymbiont. We assessed that Glossina morsitans morsitans genome contains fractions of the wolbachia gene repertoire. Wsp, fbpA and 16S sequences have been found to be inserted into the same region of X, Y and B-cromosomes. These data confirm the horizontal transmission of this endosymbiont The genomic insertion of these sequences has been also identified on Gmm supercontigs , allowing the identifications of the rearrangements that these sequences underwent in the insertion process. In collaboration with dr Aksoy we are developing transcriptome profilings from the male reproductive tissues: testes and accessory glands in order to analyse the spermatogenesis process and the machinery components of male ejaculate. Both of these processes can be influenced by the presence of pathogens and symbionts present in male reproductive tissues leading to male sterility and female fecundity/sterility Introduction Attempts to control/eradicate Glossina species using biological methods require in-depth information about the specie specific reproductive biology which may be impacted by the presence of endosymbionts and pathogens. An important aspect of mating behavior is the number of times a female mates in the wild as this influences the effective population size and may constitute a critical factor in determining the success of control methods. On the other side the presence of a polyandric behavior in a population may impact the spreading of endosymbionts / pathogens and the sterility We have considered the relations between polyandry, migration rate and the spreading of Wolbachia in wild populations. In relation to this, we have also characterized the Wolbachia Page 43 sequences which are inserted into the Glossina genome. We are also characterising the expression profiles derived from testes and accessory glands.

Main results Polyandry and Wolbachia Polyandry is a common events in wild populations of G fuscipes fuscipes and G pallidipes .Analyses have identified the presence of wolbachia in natural populations of these species. As it has suggested that Wolbachia associated incompatibilities may promote polyandry,studies are now investigating the potential influence of wolbachia on remating. As Wolbachia infections are entertainer as a tool to drive genetically desirable phenotypes into natural populations,female mate choice and remating may also have an impact on strategies on population replacement.

Glossina morsitans morsitans contains fractions of the wolbachia gene repertoire PCR screenings,sequencing, alignments and in situ hybridization on mitotic chromosomes allowed us to asses that Wsp fbpA abd 16S sequences are inserted into X,Y and B chromosomes of Glossina morsitans morsitans. . These data confirm the horizontal transfer of wolbachia. The genomic insertion of these sequences has been also identified on Gmm supercontigs, allowing the identifications of the rearrangements that these sequences underwent in the insertion process. The locations of these insertions in the sexual chromosomes suggests that X,Y and B chromospomes share a common evolutionary origin.

Development of transcriptome profiles from testes and accessory glands of Glossina morsitans morsitans We are using deep RNA seq in order to identify genes espressed in testes and male accessory glands. In order to understand the species specific processes which underlie and control spermatogenesis, mating and fertilization success. This analyses is the premise to understand the effect of wolbachia and virus on the sterilty

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4. Logical framework Narrative Summary Objective Verifiable Means of Verification Important Indicators Assumptions Overall Objective N/A N/A Continued need and Understanding and Member States exploiting interactions requests for between tsetse flies and expansion of tsetse their microbes to enhance control measures the efficacy and implementation of tsetse programmes with an SIT component Specific objectives 1. To clarify tsetse 1a. Role of symbionts 1. Publication of results. 1. Two workshops symbiont and other and other microbes in held 1) to assure microbe interactions in host physiology appropriate laboratory and natural identified sampling of field populations populations and standard DNA preparation and 2) to genotype symbionts and pathogens

1b. Genetic differences Publication of results 1b. Tsetse material within host/symbiont available for populations defined members

1c. Host/symbiont 1c. Risk analysis population genetics data maps available integrated into tsetse databases

1d. Transmission modes Publication of results of tsetse symbionts determined

1e. Compatibility of 1e. Availability of fungus infected flies with irradiated insects SIT determined

2. To understand and 2a. DNA database for 2a. DNA sequence 2. Virus-free manage tsetse virus virus populations submitted to public colonies can be interactions in laboratory identified databases developed populations 2b. Transmission mode 2b.Publication of results of tsetse SGHV determined

2c. SGH symptom free 2c. Protocols established colonies maintained and available to maintain

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colonies

3. To manipulate microbial flora to express parasite 3a. Expression system 3. Publication of results 3. Fitness of refractoriness traits enhanced resistant lines uncompromised 3b.Trypanosome Publication of results inhibitory products identified

3c. Parasite resistant lines developed 4. To improve tsetse suppression technology 4. Viable fungal 4. Effective suppression 4. Tsetse can be pathogens found on of field populations contaminated tsetse flies in the field through mating 5. To harness symbiont mediated natural mating 5a. Role of Wolbachia 5. Publication of results 5. Wolbachia incompatibilities infections in tsetse infected, physiology determined superinfected and uninfected tsetse 5b. Natural and novel CI Publication of results lines can be mating types identified. developed

5c. CI properties of tsetse Publication of results Tsetse Wolbachia Wolbachia determined transinfections can be accomplished 5d. Correlation between Wolbachia infections and Publication of results polyandry investigated

6. To disseminate 6a. Workshops knowledge among completed 6. Agency report 6. Additional disease endemic country funding for researchers to improve 6b. Partnership in PAAT, Website connection participation in field application of SIT IGGI and PATTEC established PAAT meetings and through better decision established for preparing PAAT making and capacity Creation of link(s) with position paper building 6c. Endemic country IGGI and other related available scientists trained. initiatives

PAAT position paper produced. New article for PLoS Neglected Diseases Outputs 1. Decipher host- 1. Critical symbiont 1. Functional 1. Biological symbiont interactions to products identified for relationships published material available understand tsetse’s host fitness nutritional ecology and 1b. Colonies established improve mass rearing procedures

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2. Investigate tsetse 2. Role of fungus 2. Data published 2. Availability of pathogen interactions to infections and field sites and improve control dissemination efficiency irradiated material. strategies determined

3. Study the population 3a. Wolbachia and 3. Data published 3. Availability of dynamics of tsetse Sodalis genotypes Data published field material and microbial flora identified necessary import permits 3b. Temporal and spatial analysis of microbe infection prevalence established

4. SGHV symptom-free lines established 4. Agency report Data published

4. Characterize salivary gland hyperplasic virus (SGHV) group and achieve its control in colony rearing process 5. Effectors identified 5. Data published 5. Availability of material 6. Lines to be developed 5. Identify trypanosome 6. Data published 6. Transmission inhibitory products mode known and fertility preserved 6. Develop parasite 7. Trypanosome resistant refractory lines fly lines to be produced 7. Data published 7. Selection of resistant lines possible 7. Incorporate parasite 8. Drive system resistance traits into SIT evaluated Data published 8. Wolbachia lines infected and uninfected material 8. CI mediated gene available drive system Tsetse Wolbachia association expresses high levels of CI 9. Cytoplasmic incompatibility mediated 9. Data published 9. Tsetse Wolbachia sterility confirmed association expresses high levels of CI 9. Harness cytoplasmic incompatibility for SIT 10. Workshops held application Publication of results in 10. Workshops scientific journals funded

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10. Disseminate discoveries to endemic 11. Papers published countries and interested 11. Papers accepted parties

11. Publish results in scientific journals

Activities 1. Form a network of 1. Research Contracts 1. Approval of Contracts 1. Suitable proposals research collaborators and Research and agreements by PCC- submitted and Agreements awarded NA sub-committee funding available

2. Inform PAAT and 2. CRP Information on 2. Responses to 2. Relevance of CRP PATTEC as a means to PAAT-Link participate in CRP to tsetse community disseminate future results.

3. Issue a Technical 3. Tsetse material 3. Acknowledgement by 3. Funds and flies Contract to enable supplied to CRP CRP participants available dissemination of tsetse participants fly laboratory material to CRP members

4. Issue a Technical 4. Field samples supplied 4. Acknowledgement by 4. Funds available Contract for to CRP participants CRP participants and fly populations collection/analysis and accessible dissemination of field samples to CRP members

5. Establish data base 5. Data base created 5. Open access to data 5. Resources repository base available in Seibersdorf

6. Organize 1 st RCM 6. 1 st RCM and workshop 6. Participants and CRP 6. Molecular and held in 2007 Progress Report genetic techniques appropriate

7. Organize 2 nd RCM to 7. 2 nd RCM held in 2009 7. Participants and CRP 7. Progress review molecular and and workshop held Progress Report. satisfactory genetic outputs together with a workshop to disseminate protocols 8. Organize 3 rd RCM to 8. 3 rd RCM held in 2010 8. Participants and CRP 8. Progress review molecular and and workshop held Progress Report. satisfactory genetic outputs together with a workshop to disseminate protocols

9. Organize 4 th RCM to 9. 4 th RCM held in 2012 9. Participants and CRP 9. Progress Page 49 review molecular and final Report. satisfactory genetic outputs

10. Collate all reports 10. Publish results in 10. Scientific 10. Final reports are and synthesize results refereed journal publications submitted to the Agency

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5. Agenda FOURTH RESEARCH CO-ORDINATION MEETING

JOINT FAO/IAEA DIVISION OF NUCLEAR TECHNIQUES IN FOOD AND AGRICULTURE

“Improving SIT for Tsetse Flies through Research on their Symbionts and Pathogens”

Vienna, Austria

26-30 March, 2012. VIC, ROOM M6

AGENDA

Monday, 26 March , 2012

SESSION 1

08.00-09. 00 Registration and coffee

09.00-09.15 Jorge Hendrichs and Marc Vreysen : Welcome and introduction

09.15-09.30 Administrative details

09.30-10.00 A. Abd -Alla : Glossina pallidipes Salivary gland hypertrophy virus activities in Seibersdorf

10.00-10.15 COFFEE

SESSION 2

10.15-10.45 Henry M. Kariithi. Proteomic approaches for Salivary Gland Hypertrophy Virus control in tsetse fly colonies.

10.45-11.15 Just M. Vlak. Genetic conservation of Salivary Gland Hypertrophy Virus in Africa.

11.15-11.45 Laura Guerra, John G. Stoffolano Jr, Gabriella Gambellini, Valentina Laghezza Masci, Maria Cristina Belardinelli and Anna Maria Fausto: Ultrastructure of the Salivary Glands of Non-Infected and Infected Glands in Glossina pallidipes by the Salivary Glands Hypertrophy Virus (SGHV)

11.45-12.15 Orlov Igor: The structural features of the Salivary Gland Hypertrophy Virus of the tsetse fly revealed by cryo-electron microscopy.

12.15-13.00 LUNCH

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13.00-13.30 F. Wamwiri, R. Changasi and Sam Guya: Trypanosome and SGHV co- infection in G.pallidipes and G.austeni in Kenya

13.30-14.00 Johannes A. Jehle , Yongjie Wang and Adly M. M. Abd Alla: Evolution, Phylogeny and Taxonomy of Hytrosaviridae

14.00-14.30 Basil Arif: Attempts to replicate GpSGHV in established insect cell lines

14.30-14.45 COFFEE

14.45-15.15 John P. Burand: RNAi as a Tool to Control Insect Pathogens and Parasites

15.15-15.45 Zhihong (Rose) Hu: Generation of polyclonal antibodies against hytrosaviruses

15.45-16.15 Verena Lietze : Maintenance and transmission of the disease in house fly populations

16.15-16.45 C. J. Geden, A. G. Maruniak, V.-U. Lietze, J. Maruniak, and D. G. Boucias: House fly SGHV: Impact of house fly hytrosavirus on stable fly

16.45-17.15 N. K. Maniania: The use of entomopathogenic fungi in the control of tsetse flies.

17.15-18.00 General discussion

Tuesday, 27 March , 2012

SESSION 3

08.30-09.00 Ouédraogo / Sanon G.M. Sophie : Testse flies dynamics, distribution and biology in west of Africa

09.00-09.30 Drion Boucias: Comparative Analysis of the Pathobiology of the Glossina pallipides and Musca domestica Hytrosaviruses

09.30-10.00 O. Koekemoer : Genetic characterization of Wolbachia symbionts from G. brevipalpis and G. austeni from southern Africa.

10.00-10.30 COFFEE

10.30-11.00 Vangelis Doudoumis, George Tsiamis, Florence Wamwiri, Corey Brelsfoard, Uzma Alam, Emre Aksoy, Stelios Dalaperas, Alexander P. Egyir-Yawson, Imna Malele, Johnson Ouma, Peter Takac, Adly Abd- Alla, Serap Aksoy and Kostas Bourtzis : Tsetse-Wolbachia Symbiosis: from fundamental research to novel symbiont-based pest and disease control Page 53

11.00-11.30 Wolfgang J. Miller and Daniela Schneider: Hidden Wolbachia complexity and titre dynamics in tsetse flies and their subspecies hybrids.

11.30-12.00 Malele , I.: Prevalence of SGHV among tsetse species of economic importance and their implication for SIT application

12.00-12.30 Séverine Balmand, Claudia Lohs, Serap Aksoy , Abdelaziz Heddi : Tissue distribution and transmission routes for the Tsetse fly endosymbionts

12.30-13.30 LUNCH

13.30-14.00 S. Aksoy : Tsetse symbiont and pathogen density dynamics.

14.00-14.30 Falchetto M, Malele, Bourzis K, Aksoy S and Malacrida A : Reproduction and endosimbionts in Glossina

14.30-15.00 Flobert Njiokou, : Relation between the presence of Sodalis glossinidus and the prevalence of trypanosomes in Glossina palpalis palpalis populations in Cameroon

15.00-15.30 COFFEE

15.30-16.00 F. Njiokou, O. Farikou, G. Cuny, A. Geiger .: Sodalis diversity in Cameroon

16.00-16.30 Jan Van Den Abbeele: Expression and extracellular release of a functional anti-trypanosome Nanobody in Sodalis glossinidius

16.30-17.00 Peter Takac : The determination of appropriate artificial diets to study endosymbiont-free Glossina morsitans morsitans flies

17.00-17.30 General discussion

Wednesday 28 March , 2012

SESSION 4

08.30-09.45 General Discussion of the Logical Framework and CRP evaluation documents and Formation of Two Working Groups (see below)

09.45-10.15 COFFEE

10.15-11.15 General Discussion of the Logical Framework and Formation of Two Working Groups (see below)

11.15-12.00 Working Group Discussions ( Group 1 Room M6, Group 2 ROOM MOE70

12.00-19.00 LUNCH

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13.30-17.30 Excursion organized by Peter Takac to Bratislava

Thursday 29 March , 2012

SESSION 5

09.00-09.40 Working Group Discussions

09.45-10.15 COFFEE

10.15-12.30 Working Group Discussions

12.30-13.30 LUNCH

13.30-15.30 Drafting Report

15.30-16.00 COFFEE

16.00-17.00 Drafting Report

18.00 Group Dinner (Melker Stiftskeller, Schottengasse 3, A-1010 Wien http://www.melkerstiftskeller.at/index.php?option=com_content&view= article&id=50&Itemid=59&lang=en

Friday 30 March , 2012

SESSION 6

09.00-10.30 Reports of Working Groups and CRP evaluation

10.30-11.00 COFFEE

11.00-12.30 Drafting of CRP evaluation

12.30-13.30 LUNCH

14.00-14.30 General Discussion

Closing

Working Group 1: Symbionts (Room M6) Aksoy, Wolfgang, Bourtzis, Geden, Heddi, Njiokou, Malacrida, Koekemoer, Egyir- Yawson, Takac, Geiger, Jan, Franco, Daniela

Working Group 2: Pathogens (Room MoE70) Boucias , Vlak, Jehle, Bergoin, Abila, Malele, Ouedraogo, Wamwiri, Maniania, Basil, John, Laura, Orlov, Hu, Patrick. . Page 55

6- LIST OF PARICIPANTS LIST OF PARTICIPANTS Final RCM on “Improving SIT for Tsetse Flies through Research on their Symbionts and Pathogens” 26-30 March 2012 Vienna, Austria AUSTRIA Sault Ste. Marie Mr Wolfgang Miller ON P6A 2E5 Laboratories of Genome Dynamics Tel: 001 705 541 5512 Medizinische Universität Wien Fax: 001 705 541 5700 Währingerstrasse 10 Email: [email protected] 1090 Wien Tel: 0043 1 4277 60625 CHINA Fax: 0043 1 4277 60690 Ms Zhihong Hu Email: [email protected] Wuhan Institute of Virology Chinese Academy of Sciences BELGIUM Xiaohongshan 44 Mr J. Van Den Abbeele Wuhan 430071 Prince Leopold Institute of Tropical Medicine Tel: 0086 27 87197180 Nationalestraat 155 Fax: 0086 27 87197180 2000 Antwerpen Email: [email protected] Tel: 32 3 2476311 Fax: 32 3 247 6359 FRANCE Email: [email protected] Mr Max Bergoin Université Montpellier II BURKINA FASO Place Eugène Bataillon Ms Gisele M. S. Ouedraogo 34095 Montpellier CEDEX 05 Laboratoire National D’Elevage Tel : 0033 467 14 45 11 Direction Générale des services vétérinaires Fax : 0033 467 14 42 99 B.B. 7026 Email : [email protected] Tel : 226 50 32 46 35 Email : [email protected] Ms Anna Geiger Institut de recherche pour le dévelopement CAMEROON (IRD) Mr Flobert Njiokou 213, rue Lafayette Faculté des Sciences 75480 Paris CEDEX 10 Université de Yaounde Tel: 0033 467 59 38 35 B.P. 812 Fax: 0033 467 59 39 20 Yaoundé Email: [email protected] Tel : 237 223 44 96/237 771 96 31 [email protected] Fax : 237 223 44 96 Email : [email protected] Mr Abdelaziz Heddi UMR INRA/INSA de Lyon BF2I CANADA Bâtiment Louis Pasteur, INSA de Lyon, Mr Basil Arif 11, Avenue Jean Capelle, Laboratory for Molecular Virology 69621 Villeurbanne cedex Great Lakes Forestry Centre Tel: 0033 472 43 88 68 1219 Queen St. E. Fax: 0033 4 72 43 85 34 Email: [email protected]

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International Centre of Insect Physiology GERMANY & Ecology (ICIPE) Mr Johannes Jehle P.O. Box 30772 Head of Institute for Biological Control Nairobi Julius Kühn-Institute Tel: 00254 20 863 2000 Heinrichstr. 243 Fax: 00254 20 863 2001/2 64287 Darmstadt Email: [email protected] Tel: +49 6151 407-220 Fax: +49 6151 407-290 Ms Florence Wamwiri [email protected] Trypanosomiasis Research Institute P. O. Box 362 GHANA Muguga, Kikuyu Mr Alex Egyir Yawson Tel: 254 020 2700604/020 2700654 Biotechnology & Nuclear Agriculture Email: [email protected] Research Institute [email protected] Ghana Atomic Energy Commission (GAEC) P.O. Box 80 NETHERLANDS Legon Accra Mr Just Vlak Tel: 233 21 402286 Wageningen University Fax: 233 21 402286 P.O. Box 9101 Email: [email protected] 6700 HB Wageningen Tel: 0031 317 483090 GREECE Fax: 0031 317 484820 Mr Kostas Bourtzis Email: [email protected] Department of Environmenatl & Natural Resources Management SLOVAKIA University of Ioannina Mr Peter Takac 2 Seferi Street Institute of Zoology 30100 Agrinio Slovak Academy of Sciences Tel: 30 26410 74114 Dubravska cesta 9 Fax: 30 26410 39571 84506 Bratislava Email: [email protected] Tel: 0042 125 930 2642 Fax: 0042 125 930 2646 ITALY Email: [email protected] Ms Anna Malacrida Dipartimento di Biologia Animale SOUTH AFRICA Via Ferrata 1 Mr Otto Koekemoer Università di Pavia Agricultural Research Council (ARC) 27100 Pavia Private Bag X05 Tel/Fax +39 0382 986059 0110 Onderstepoort Email: [email protected] Tel: 27 (0) 12 5299229 Fax: 27 (0) 12 5299270 Ms Laura Guerra Email: [email protected] Universita degli Studi della Tuscia Via San Giovanni Decollato, 1 UGANDA 01100 Viterbo Mr Patrick P’Odyek Abila* Tel: 0039 0761 357389 National Livestock Resources Research Email: [email protected] Institute National Agricultural Research Organisation KENYA (NARO) Ms Nguya Kalemba Maniania P.O. Box 96, Tororo

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Tel: 256 45 44448360 Gainesville FL 3260 Email: [email protected] Tel: 00352 374 5919 [email protected] Fax: 00352 374 5922 Email: [email protected] UNITED REPUBLIC OF TANZANIA Ms Imna Issa Malele Tsetse Department OBSERVERS Tsetse and Trypanosommiasis Research USA Institute (TTRI) Ms Verena Lietze Majani Mapana University of Florida P.O. Box 1026 Gainesville, FL 32611 Tanga 053 Tel: 001 352 374 5919 Tel: 255 27 2642557/ 2644572/3 Fax: 001 352 374 5922 Fax: 255 27 2642557/ 27 2643783 Email: [email protected] Email: [email protected] [email protected] AUSTRIA Daniela Schneider UNITED STATES OF AMERICA Lab of Genome Dynamics Ms Serap Aksoy Dep. Cell and Developmental Biology Yale University Medical University of Vienna Dept. of Epidemiology & Public Health Waehringerstr. 10 New Haven, CT 06520 1090 Vienna Tel: 001 203 737 2180 Email: [email protected] Fax: 001 203 785 4782 Email: [email protected] Italy Marco Falchetto Mr John Burand University of Pavia Department of Plant Soil and Insect Sciences Department of Biology and Biotechnology Department of Microbiology Via Ferrata 9, 27100 Pavia, Italy University of Massachussetts Amherst Email: [email protected] Amherst, MA 01003 tel: +39 0382 986023 Tel: 413-545-3629 Fax 413-545-2115 The Netherland Email: [email protected] Dr. İkbal Agah İNCE, Assistant Professor Adjunct Scientist, Molecular Virology CONSULTANTS Wageningen University and Research Center Laboratory of Virology FRANCE Radix Building 107, Droevendaalsesteeg 1 Mr Igor Orlov 6708 PB Wageningen, The Netherlands Institut de Génétique, de Biologie Moléculaire et Cellulaire Tel: 0031317483099 B.P. 10142 Fax: 0031317484820 67404 Illkirch Cedex Email: [email protected] Email: [email protected] IAEA Mr Henry Kariithi UNITED STATES OF AMERICA Joint FAO/IAEA Division Mr Christopher Geden USDA Agriculture Research Service Center Insect Pest Control Laboratory For Medical Agricultural & Veterinary Seibersdorf [email protected] Entomology 1600 SW 23 rd, Email:

* Participants did not attend the 4th RCM in Vienna

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ANNEX I: WORKING PAPERS Abdelaziz Heddi: Tissue distribution and transmission routes for the Tsetse fly 61 endosymbionts. (Anne Geiger) Njiokou F., Farikou O., Cuny G. 2 & Geiger A.: Sodalis diversity in 65 Cameroon Basil Arif: Attempts to replicate GpSGHV and MdSGHV in tissue culture. 73 Christopher J. Geden: Final report, MdSGHV research conducted at the USDA lab in 77 Gainesville, Florida Drion Boucias, Vereena Lietze. Christopher Geden, and Zhang Hu: Studies on the 81 Hytrosavirus infecting the house fly Musca domestica Flobert Njiokou. Final report on the IAEA Research Contract N° 14320. 85 Florence Wamwiri, Samuel Guya and Robert Changasi: Investigations on the 91 presence and effects of Wolbachia and the salivary gland virus (SGHV) in laboratory and wild populations of Glossina pallidipes and G. austeni. Gisele Oudraogo, Anicet Georges Ouédraogo, Issa Sidibé, Jean Baptiste Rayaissé, 95 Adly M.M. Abd-Alla,: Testes flies dynamics, distribution and biology in west of Africa. Imna Malele,I.: : Prevalence of salivary gland hypertrophy virus (SGHV) in different 111 tsetse populations and the possibility of establishing SGHV free tsetse colonies Jan Van Den Abbeele, Linda De Vooght,& Guy Caljon: Expression and extracellular 123 release of a functional anti-trypanosome Nanobody® in Sodalis glossinidius, a bacterial symbiont of the tsetse fly. Johannes A. Jehle, : Final report 129 John P. Burand, Justin Nguyen and Woojin Kim: RNAi Inhibition of HzNV-2 131 Pathology in Helicoverpa zea Moths. Just M. Vlak, Henry M. Kariithi, Jan W.M. van Lent, Adly M. M. Abd-Alla, Monique 139 M. van Oers and Just M. Vlak: Structure, protein composition, morphogenesis and cytopathology of Glossina pallidipes salivary gland hypertrophy virus to develop rationally-designed immuno-intervention strategies to prevent virus transmission in tsetse fly colonies. Kostas Bourtzis, Vangelis Doudoumis, George Tsiamis, Florence Wamwiri, Corey 143 Brelsfoard, Uzma Alam, Emre Aksoy, Stelios Dalaperas, Alexander P. Egyir- Yawson, Imna Malele, Johnson Ouma, Peter Takac, Adly Abd-Alla, Serap Aksoy: Tsetse-Wolbachia Symbiosis: from fundamental research to novel symbiont-based pest and disease control strategies. Laura Guerra, John G. Stoffolano Jr, Gabriella Gambellini, Valentina Laghezza 175 Masci, Maria Cristina Belardinelli and Anna Maria Fausto : Ultrastructure of the Salivary Glands of Non-Infected and Infected Glands in Glossina pallidipes by the Salivary Glands Hypertrophy Virus (SGHV) Max Bergoin and François Cousserans: Final report. 193 Nguya K. Maniania: Understanding and exploiting interactions between tsetse flies 197 and their microbes to enhance the efficacy and implementation of tsetse. programmes with an SIT component. Otto. Koekemoer: The role of pathogens and symbionts of Glossina brevipalpis and G. 205 austeni in South Africa . Peter TAKAC : The efect of antibiotic and yeast treatment on Glossina genus tsetse 215

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flies (Rose) Zhihong Hu: Generation and evaluation of polyclonal antibodies against 219 hytrosaviruses Serap. Aksoy: Final report. 223 Wolfgang J. Miller and Daniela Schneider: Evaluating Wolbachia prevalence, strain- 227 fingerprinting tools, and experimental hybrid formation in Glossina

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Title Tissue distribution and transmission routes for the Tsetse fly endosymbionts

Authors

Séverine Balmand 1,2 , Claudia Lohs 3, Serap Aksoy 3, Abdelaziz Heddi 1,2

1INSA-Lyon, INRA, UMR203 BF2I, Biologie Fonctionnelle Insectes et Interactions, F- 69621 Villeurbanne, France, 2Université de Lyon, F-69003, Lyon, France; 3Yale University School of Public Health, LEPH 816, 60 College Street, New Haven, Connecticut 06520, USA.

Abstract During this CRP program, we have studied the precise tissue distribution of tsetse endosymbiotic bacteria and have infered their respective way of transmission from parents to progenies. To this end, we conducted a Fluorescence In situ Hybridization (FISH) study to survey bacterial spatial distribution across the fly tissues. We showed that bacteriocytes are mono-infected with Wigglesworthia , while both Wigglesworthia and Sodalis are present in the milk gland lumen. Sodalis was further seen in the uterus, spermatheca, fat body, milk and intracellular in the milk gland cells. Contrary to Wigglesworthia and Sodalis, Wolbachia were the only bacteria infecting oocytes, trophocytes, and embryos at early embryonic stages. Furthermore, Wolbachia were not seen in the milk gland and in the fat body. This work further highlights the diversity of symbiont interactions in multipartner associations and supports two maternal routes of symbiont inheritance in the tsetse fly: Wolbachia through oocytes, and, Wigglesworthia and Sodalis by means of milk gland bacterial infection at early post-embryonic stages.

Introduction The tsetse fly Glossina is the vector of the protozoan Trypanosoma brucei spp., that causes Human and Animal African Trypanosomiasis in sub-Saharan African countries. To supplement their unbalanced vertebrate bloodmeal diet, flies permanently harbor the obligate bacterium Wigglesworthia glossinidia, which resides in bacteriocytes in the midgut bacteriome organ as well as in milk gland organ. Tsetse flies also harbor the secondary facultative endosymbionts (S-symbiont) Sodalis glossinidius that infects various tissues and Wolbachia that infects germ cells. However, while the tsetse symbiont functions are being deciphered, less is known about the precise tissue localization and the way by which each symbiotic partner is transmitted from parent to offspring. The tsetse reproduction is unique among insects as it is viviparous and females carry and nourish their progeny during the entire larval development. A single oocyte develops at a time. After ovulation and fertilization, the embryo hatches within the uterus, while a second oocyte begins its development in the second ovary. During the intrauterine development, the mother nourishes the offspring by milk produced by differentiated accessory gland (named the milk gland), which empties into the uterus. Therefore, the progeny can acquire the symbiont infections either through germ line cells, or through milk secretions of the mother at early larval stages during their maturation in the uterus. The aim of this work was to understand how tsetse endosymbiotic bacteria are

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transmitted from one generation to the other, by analyzing the precise localization of the endosymbionts in host tissues associated with reproduction. To this end, we have used Fluorescent In Situ Hybridization (FISH) by using specific fluorescent oligoprobes designed to match specifically with 16S rRNA of the different bacteria.

Main results To better understand how tsetse endosymbionts are transmitted from the mother to her progeny, we have examined the symbiont distribution across mother’s tissues involved in reproduction ( i.e. the reproductive tract and tissues nourishing developing progeny), and within the progeny at early developmental stages. To this end, we have used FISH experiments with probes designed to match specifically with the 16S rRNA of each endosymbiont to avoid any cross-hybridization. All probes were labeled with a fluorescent tail so that bacteria can be analyzed by fluorescence microscopy. Probe specificity was tested by hybridizing each probe with tissues housing separately Wigglesworthia , Sodalis and Wolbachia . Figure 1 shows, for example, the specificity of Sodalis , and Wolbachia probes with regards to Wigglesworthia. Table I summarizes the samples analyzed in this study and the presence/absence of each symbiont across insect tissues and organs.

1. The bacteriome tissue houses Wigglesworthia only As an obligate endosymbiont, Wigglesworthia is contained in bacteria-bearing host cells called the bacteriocytes, which are grouped to form the bacteriome organ. This tissue is located in the anterior midgut forming two lobes surrounding the gut. FISH experiments with the Wigglesworthia probe showed strong signals from bacteriome sections from the female gut. Each bacteriome lobe is formed by different bacteriocyte layers (Balmand et al., 2012 JIP). Remarkably, applying probes of S-symbionts failed to show any signal on the same sections, suggesting a monoinfection status of tsetse bacteriocytes with the primary endosymbiont (P-symbiont) Wigglesworthia.

2. Wigglesworthia does not infect reproductive tissues FISH analysis of the female reproductive tract with Wigglesworthia probe failed to reveal any signal from ovaries and uterus (Balmand et al., 2012 JIP). Surrounding adipocytes were also free of Wigglesworthia . Accordingly, the examination of 194 oocytes, trophocytes and embryos did not show any Wigglesworthia inside these germ cells. However, Wigglesworthia was abundant in the lumen of the mother’s milk gland tubules, while some bacterial cells were also observed in the canal leading to the secretory cell vacuoles, where the milk products are produced and secreted to feed the developing progeny in the uterus. Wigglesworthia could also be found extracellularly in cavities within the fat body in larva. As for the progeny, infection with Wigglesworthia seems to occur late during development. Wigglesworthia was detected in the first instar larvae, at low amounts surrounding the intestine. The bacterial density increased in the bacteriome of the third instar larvae. Interestingly, Wigglesworthia does not infect the larval intestine but remains confined to the second layer of cells that surround the gut making up the bacteriome, similar to that described in Blochmannia association with the carpenter ant Camponotus floridanus . Taken together, these data indicate that Wigglesworthia constitutes two separate sub-populations in adult females: the first strictly intracellular within the bacteriocytes in the adult gut and the second free-living in the lumen of milk glands and inside fat body cavities in larva.

3. Sodalis endosymbionts are in the reproductive tract and infect milk gland cells and

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lumen. As was the case for Wigglesworthia, no Sodalis cells were detected in oocytes and embryos (Balmand et al., 2012 JIP). However, unlike Wigglesworthia , Sodalis cells are spread throughout the reproductive tract. They were shown to infect the oviduct, all over the uterine muscle tissue, in the fat body, and even in spermatheacae sheath cells. Moreover, Sodalis can be detected in the lumen of the milk gland , intracellular within the secretory cells, or in both compartments. In the milk gland lumen, cohabitation between Sodalis and Wigglesworthia was shown is only few cases, indicating a possible competition between these bacteria. In young larval progeny, the S-symbiont Sodalis can be detected in the first instar larvae, as the P-symbiont Wigglesworthia . However, while Wigglesworthia invades the bacteriocyte cells and differentiates into the bacteriome organ, Sodalis cells form aggregate islands throughout the fat body and gut.

4. Wolbachia infects early oocytes and embryos Wolbachia endosymbiont is the only bacterium that infects embryos prior to intrauterine development. Wolbachia was seen in the developing oocytes, being at both poles as a thin layer, within the yolk, just above the follicular cells. In intimate connection with the oocytes, four out of five trophocytes analyzed were highly infected with Wolbachia . Apart from these cells, Wolbachia was not detected in the fat body and in the uterus. Trophocytes could represent a Wolbachia reservoir, intimately connected to oocytes. When oocytes move down through the uterus to hatch into an embryo, Wolbachia can still be detected at both poles of the embryo, constituting aggregates on the follicular cell layer. Later in second instar larvae, Wolbachia was seen sporadically in aggregates. Contrary to the nutritional endosymbionts Wigglesworthia and Sodalis , Wolbachia was not detected in the milk gland. It is unlikely that milk secretions provide a route of transmission for Wolbachia .

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Sodalis diversity in Cameroon

Njiokou F. 1, Farikou O. 1, Cuny G. 2, Geiger A. 2

1 General Biology Laboratory, University of Yaoundé I, Faculty of Science, BP 812 Yaoundé, Cameroon 2 UMR 177, IRD-CIRAD, CIRAD TA A-17/G, Campus International de Baillarguet, 34398 Montpellier Cedex 5, France

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Introduction In a previous work, a large tsetse fly sampling campaign had been conducted in Bipindi and Campo that are two historical sleeping sickness foci located in the south of Cameroon. The results evidenced large differences between the population of flies from Campo and that from Bipindi. So we hypothesized that the geographical isolation of the two foci could have induced independent evolution of fly, and symbiont populations, leading to a divergent diversification of genotypes. To test this hypothesis, we investigated the symbiont genetic structure using the allelic size variation at four specific microsatellite loci. The results showed that the genetic structure of samples varied at different geographical scales, with almost no differentiation among the villages of the Campo Human African Trypanosomiasis (HAT) focus and a low but significant differentiation between the Campo and Bipindi HAT foci. So, the data provided new information on the genetic diversity of S. glossinidius populations. Possible interactions between S. glossinidius subpopulations and Glossina species that could favor tsetse fly infections by a given trypanosome species should be further investigated.

Materials and Methods Collection of Sodalis glossinidius Glossina palpalis palpalis flies were collected in two HAT foci (Bipindi and Campo) situated in the Ocean Division of the South Region of Cameroon. The Campo focus (2°20 ′N, 9°52 ′E) presents several biotopes (farmland, marshes, swampy areas, and equatorial forest). The Bipindi focus (3°2 ′N, 10°22 ′E) has a typical forest bioecological environment including equatorial forest and farmland along roads and around villages. Entomological surveys were conducted in 2007 in Bipindi and Campo. The geographical positions of the sampling sites were determined using the global positioning system. Tsetse flies were captured using pyramidal traps planted in suitable tsetse fly biotopes. Each trap remained deployed for four consecutive days and flies were harvested twice a day. The different Glossina species were first identified and then sorted into teneral and non-teneral flies, according to morphological criteria. Flies were dissected in a drop of sterile 0.9% saline solution, and their midgut separately transferred into microfuge tubes containing ethanol (95°) for further symbiont analyses.

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DNA extraction Total genomic DNA was extracted from the midguts using a standard protocol. Briefly, the material was homogenized in CTAB Buffer (CTAB 2% ; 0,1M Tris pH8 ; 0,02M EDTA pH8; 1,4M NaCl) with a pestle and incubated at 60° C for 30 min. DNA was extracted by chloroform / isoamilic alcohol, isopropanol precipitated and resuspended in water.

Selection of microsatellites in the Sodalis glossinidius genome and DNA amplification We used four microsatellites described in Farikou et al., 2011a; 2011b. Midguts showing specific detection of S. glossinidius were further processed for S. glossinidius microsatellite genotyping. The method was adapted from Farikou et al., 2011a.

Electrophoresis of PCR products After specific amplification, infrared dye-labeled (IRD700 or IRD 800) PCR products were diluted to 1/5, 1/10, or 1/50 in loading buffer (95% deionized formamide, 20 mM ethylenediaminetetraacetic acid (EDTA), pH 8.0, and 1 mg/ml bromophenol blue). Then they were denatured for 3 min at 95°C, and transferred to ice before loading. The sample-loading volume was 1.22 µl. Each mixture was separated, in a 1- to 2-h run at 1500 V, on a 6.5% (wt/vol) Long Ranger polyacrylamide gel, using 1 x Tris-borate-EDTA buffer, on a two-dye, model 4300 LI-COR-automated DNA sequencer. Infrared images of the patterns were analyzed using the semiautomated scoring program Quantar. Measurement of allele length on polyacrylamide gels was automated using molecular size markers.

Statistical analyses Genetic diversity estimation Finally, genetic diversity was analyzed on 244 samples carrying the symbiont. Of these, 131 were from the Campo focus, corresponding to three villages: 38 from Akak village (N 02°22.831', E 09°58.654'), 33 from Campo Beach/Ipono (N 02°20.985', E 09°50.300'), and 60 from Mabiogo (N 02°17.657', E 09°51.938'). 113 samples were from the Bipindi focus: Ebimimbang village (N 03°02.856', E 10°28.515'). The populations corresponded to the four villages. The mean number of alleles per locus, the allele frequencies, and the level of heterozygosity ( HE) were estimated for each locus and each population using ARLEQUIN software, version 3.5.1. Microsatellite alleles were then combined into haplotypes to perform the following analyses. Each population was

Page 68 characterized by its level of diversity using the number of detected haplotypes, the haplotypic diversity ( HEh ), and the rarefied haplotypic richness ( HR), computed using the CONTRIB 1.02 program. The rarefied haplotypic richness ( HR) is defined as the expected number of different haplotypes found in each population using a standardized sample size fixed as the smallest available number of genotyped individuals.

Genetic structure The genetic structure among S. glossinidius populations, at the village level and the HAT foci, was tested using the analysis of molecular variance (AMOVA, ARLEQUIN) based on haplotype frequencies. AMOVA subdivided the genetic diversity into hierarchical components and estimated three fixation indices: FCT , which can be interpreted as the relative divergence between HAT foci (Bipindi and Campo foci), FSC , corresponding to the relative divergence between populations within the Campo focus, and FST , corresponding to the relative divergence among populations. FCT was tested by permuting populations among foci,

FSC by permuting haplotypes among populations within a focus, and FST by permuting haplotypes among populations among the foci. Moreover, pairwise FST between populations were computed and their significance was tested by 10,000 permutations using ARLEQUIN. To better understand the molecular relationships between intraspecific data, we connected haplotypes using a median-joining network with the NETWORK 4.5.1.6. program, equally weighting each locus, and setting the epsilon parameter to 20 to obtain a full median network. The median-joining network algorithm combines the Kruskal algorithm for finding minimum spanning trees and Farris’s maximum-parsimony heuristic algorithm.

Geospatial analysis using GenGIS GenGIS was used to visualize haplotype diversity and its relationship between geographically distant populations.

Results Genetic diversity in S. glossinidius The complete dataset included multilocus genotypes for the 244 S. glossinidius strains from the 244 G. palpalis palpalis sampled in HAT foci in Cameroon (113 from Bipindi, 131 from Campo). The genome of these 244 S. glossinidius samples carried the four loci investigated in

Page 69 full length. The four microsatellite loci were polymorphic, and a total of 19 alleles were detected, ranging from three (ADNg 12/13) to six (ADNg 21/22, ADNg 15/16) alleles per locus. The mean theoretical heterozygosity ( HE) was quite different between loci, ranging from 0.07 (ADNg 12/13) to 0.59 (ADNg 15/16). Per population over the four loci, the theoretical heterozygosity ( HE) varied from 0.35 to 0.41, corresponding to the villages Ebimimbang and Campo Beach/Ipono, respectively. The combination of the microsatellite alleles yielded a total of 35 haplotypes. The four populations were polymorphic, showing 12-20 haplotypes.

Population differentiation patterns The population structure of S. glossinidius was explored at different hierarchical levels using AMOVA. On haplotypic frequencies, AMOVA revealed that most of the variation was found among individuals within populations (97.8%). The fixation index reflecting the nested design of the samples indicated no overall differentiation between populations within the Campo focus ( FSC =0.003, P=0.33) and a slight but not significant differentiation at the foci level

(FCT =0.019, P=0.25). The genetic differentiation among the four populations was low

(FST =0.022) but significant ( P=0.006).

No differentiation was shown between the villages Akak and Campo Beach/Ipono ( FST =-

0.005) nor between Campo Beach/Ipono and Mabiogo ( FST =-0.001). A positive but not significant differentiation ( FST =0.009) was recorded between Mabiogo and Akak. Even though FST values were relatively low, significant differences were shown between

Ebimimbang and Mabiogo, Akak and Campo Beach/Ipono, with FST of 0.018 ( P=0.023), 0.022 ( P=0.027), 0.032 ( P=0.014), respectively. This indicates a significant differentiation between the Bipindi (Ebimimbang) population and those of the three villages (Akak, Campo Beach/Ipono, and Mabiogo) in the Campo focus.

Haplotype network and distribution The four main haplotypes (H11, H14, H27, H30), showing overall frequencies above 0.05, were shared by the populations from the four sampling areas and were present at high frequencies within the populations studied (except H27 in Ebimimbang, present at a low frequency). The median-joining network resulted in a complex haplotype network and did not show a clear pattern of phylogeographic evolution. In addition, GenGIS was used to draw a georeferenced pattern of haplotype diversity. Haplotypes with high frequencies are shared between populations. Moreover, the Akak and

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Campo Beach/Ipono populations displayed more haplotypes with frequencies above 0.05 (eight and five haplotypes, respectively) than Mabiogo (four haplotypes) and Ebimimbang (four haplotypes), reflecting the large genetic diversity of the first two populations.

Discussion The present study was conducted within our investigations on sleeping sickness and most particularly on the tripartite interactions between the vector (the tsetse fly), its secondary symbiont ( Sodalis glossinidius ), and the parasite (the trypanosomes) the vector transmits to humans and animals. This study was the first to take an interest in the population genetics of S. glossinidius in the field. Genetic diversity was lower in Mabiogo and Ebimimbang than in Akak and Campo Beach/Ipono. The lower genetic diversity of the Ebimimbang and Mabiogo S. glossinidius population may be associated with a lower effective population size in these villages. This could be due to the lower effective population size of its host, G. palpalis palpalis , or to the existence of a selective pressure exerted by the tsetse flies on the symbiont S. glossinidius in the populations concerned. Within the Campo HAT focus, differentiation between populations was not significant. The village of Ebimimbang, located in the Bipindi HAT focus, showed significant FST with the three villages in the Campo focus, with the foci 150 km apart and located on different river basins. The differentiation analysis, based on the pairwise FST between populations and the AMOVA, revealed that the S. glossinidius populations presented a slight but significant differentiation between the Bipindi and Campo HAT foci. The network and the georeferenced haplotype analysis showed that three frequent ancestral haplotypes were shared between the four populations and that there was not a geographic pattern of haplotypic diversity. These data suggest either that the gene exchange between populations occurred repeatedly or that the haplotypes derived from a common ancestral population. As a symbiont of Glossina , with mainly a vertical transmission but also perhaps a horizontal transmission among matrilines of tsetse flies, the genetic diversity of S. glossinidius depends on its host. Our results suggest that gene flow exists between tsetse flies within the Campo HAT focus and that structuring may exist between the two foci, implying a limited gene flow, at least of female flies. The slight local differentiation among the S. glossinidius populations might be related to the fly migration rate between the HAT foci. The two HAT foci are

Page 71 located on different river basins, but tsetse flies could move from place to place and form a continuous belt, which could be promoted by the presence of a large number of rivers and stream habitats, combined with suitable host availability allowing good dispersal conditions and a less confined spatial distribution of flies. In conclusion, these results provide new information on the genetic diversity of S. glossinidius populations. They evidence the existence of differences between symbiont populations according to the flies' origin, the Campo or the Bipindi HAT focus. The evidence of a slight gene flow (or gene flow maintained up to very recently) between the two foci located about 150 km from each other was unexpected. This means that tsetse fly migration occurs despite this rather large distance. This finding is important in the context of sustainable vector control.

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Attempts to replicate GpSGHV and MdSGHV in tissue culture

Basil Arif Laboratory for Molecular Virology, Great Lakes Forestry Centre, Sault Ste. Marie, Ontario, Canada

Approximately 129 cell lines from insects belonging to various insect orders have been successfully initiated and used over the years to study:

• Replication of Nucleopolyhedroviruses and Entomopoxviruses in vitro . We have looked into the replication of lepidopteran nucleopolyhedroviruses in cell lines and studied the effects of virus replication on cellular protein synthesis, isolation of specific viral ptoeins and mass propagation. Similarly, involved in the replication of entompoxviruses in different cell lines were investigated and tagged viruses were produced and used to follow the virus infection pathway in permissive larvae. • Insect physiology. We are interested in the regulation of the moulting process in the eastern spruce budworm, Choristoneura fumiferana and how the moulting events effect the replication of viruses in larvae. It is known that approximately 30 genes are transactivated when 20-hydroxyecdyson binds to the ecdysone response element in the cellular DNA. The events can be mimicked in cell lines and the expression of various genes involved in moulting has been studied. One gene, CHR3, which is a transcription factor was isolated, cloned and its value in pest control was assessed as per the next section • Generation of genetically modified viruses with enhanced capacity to control forest insect pests. A number of genetically modified NPVs against the eastern spruce budworm were constructed and their efficacy in the target animal was assessed. One of the most promising construct was a virus carrying the moulting transcription factor, CHR3, which when given to an early instar larva, it went into early moulting but part of the cuticle and the mouth parts were not tanned. This resulted in an insect unable to forage. • Effects of various bacterial toxins on insect cells. An in vitro lawn assay was developed to measure relative toxicities of various bacterial toxins to insects. This assay is very valuable in assessing the level of toxicity of various proteins to larvae. • Isolation of viruses from different insects. New cell lines have been initiated from new and emerging insect pests in Canada. One, the emerald ash borer, which particularly destructive to ash trees, appears to have a cypovirus that was isolated by inoculating a homogenate of this insect in a newly established cell line

The above studies are but a short list describing the utility of cell lines in the study of viruses, insect physiology and responses to various proteins. Certainly, they can be used to take the level of research on GpSGHV and MdSGHV into another level. One can use the cell lines to titerate these viruses, conduct molecular studies, isolate genotypic Page 74 variants, etc. We have chosen 29 cell lines from different insect orders and derived from various larval tissues to see if either SGHV virus can replicate in vitro . The selected cell lines are delineated in the table below.

CHORISTONEURA OCCIDENTALIS CO21 pupal ovaries ORGYIA LEUCOSTIGMA CO112 pupal ovaries OL4 neonate larvae CO63 neonate CO131 embryonic NEODIPRION LECONTEI

NL1 embryonic CHORISTONEURA FUMIFERANA NL18 embryonic CF203 midgut CF16 ovarian PISSODES STROBI CF70 neonate PS1 neonate

PS6 neonate LYMANTRIA DISPAR LYGUS LINEOLARIS LD8 ovarian LI21 neonate-free LD301 embryonic LI27 neonate attached LD404 neonate LI80 embryonic attached

MALACOSOMA DISSTRIA MD63 larval ovaries MD66 larval haemocytes MS5 neonate larvae

In 2011, the following 8 cell lines were resurrected from liquid nitrogen and used in attempts to replicate GpSGHV.

Lymantira dispar 1 cell line, LD652 Choristoneura fumiferana 2 midgut cell lines 1 embryonic 1 ovarian Neodiprion lecontei 2 embryonic Choristoneura occidentalis 1 neonate

Lymantria dispar cells – Ld652. An example of cells used

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Mock infected Cells Cells inoculated with GpSGHV

Each line was seeded into 48 flat well plates, allowed to attach then inoculated with either GSGHV of MSGHV and let adsorb for 1 hour. Medium (100 ul) was added to each well and the cells were examined twice daily for evidence of virus replication in comparison to mock infected cells. No evidence of virus replication was observed in the inoculated cells in terms of cells’ doubling time, cytopathic effects, occasional EM observations or amplification by PCR of a sequence in the MSGHV genome. Both the mock infected and inoculated cells appear to grow at approximately the same rate. Page 76

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Report for the workshop “Improving SIT for tsetse flies through research on symbionts and pathogens”, Vienna, Austria (March 26-30, 2012).

TITLE OF WORKING PAPER: Final report, MdSGHV research conducted at the USDA lab in Gainesville, Florida

AUTHOR: C. J. Geden

USDA, ARS, Center for Medical, Agricultural and Veterinary Entomology, 1600 SW 23 rd Dr., Gainesville, FL, USA 32608

Dr. Geden’s role in the CRP was of that of a consultant and there were no signed contractual obligations. Research was conducted on house fly SGHV in support of the CRP objective on the virus in tsetse. The following abstracts summarize three publications on this subject; additional publications with Geden as co-author are Prompiboom et al. (2010), and four papers by Lietze et al. in 2007-2012.

A survey (2005Ð2006) of house fly, Musca domestica L. (Diptera: Muscidae) populations on four Florida dairy farms demonstrated the presence of flies with acute symptoms of infection with salivary gland hypertrophy (SGH) virus on all farms. Disease incidence varied among farms (farm averages, 0.5-10.1%) throughout the year, and it showed a strong positive correlation with fly density. Infections were most common among flies that were collected in a feed barn on one of the farms, especially among flies feeding on wet brewers grains (maximum 34% SGH). No infections were observed among adult flies reared from larvae collected on the farms, nor among adults reared from larvae that had fed on macerated salivary glands from infected flies. Infected female flies produced either no or small numbers of progeny, none of which displayed SGH when they emerged as adults. Healthy flies became infected after they fed on solid food (a mixture of powdered milk, egg, and sugar) that had been contaminated by infected flies (42%) or after they were held in cages that had previously housed infected flies (38.6%). Healthy flies also became infected after they fed on samples of brewers grains (6.8%) or calf feed (2%) that were collected from areas of high fly visitation on the farms. Infection rates of field-collected flies increased from 6 to 40% when they fed exclusively on air-dried cloth strips soaked in a suspension of powdered egg and whole milk. Rates of virus deposition by infected flies on food were estimated by quantitative polymerase chain reaction at_100 million virus copies per fly per hour. Electron Page 78 microscopy revealed the presence on enveloped virus particles in the lumen of salivary glands and on the external mouthparts of infected flies. (Geden C. J., V. Lietze, and D.G. Boucias. 2008. Seasonal prevalence and transmission of salivary gland hypertrophy virus of house flies (Diptera:Muscidae). J. Med. Entomol. 45: 42-51)

House flies ( Musca domestica ) infected with Musca domestica salivary gland hypertrophy virus (MdSGHV) were found in fly populations collected from 12 out of 18 Danish livestock farms that were surveyed in 2007 and 2008. Infection rates ranged from 0.5% to 5% and averaged 1.2%. None of the stable flies ( Stomoxys calcitrans ), rat- tail maggot flies ( Eristalis tenax ) or yellow dung flies ( Scathophaga stercoraria ) collected from MdSGHV-positive farms displayed characteristic salivary gland hypertrophy (SGH). In laboratory transmission tests, SGH symptoms were not observed in stable flies, flesh flies ( Sarcophaga bullata ), black dump flies ( Hydrotaea aenescens ), or face flies ( Musca autumnalis ) that were injected with MdSGHV from Danish house flies. However, in two species (stable fly and black dump fly), virus injection resulted in suppression of ovarian development similar to that observed in infected house flies, and injection of house flies with homogenates prepared from the salivary glands or ovaries of these species resulted in MdSGHV infection of the challenged house flies. Mortality of virus-injected stable flies was the highest among the five species tested. Virulence of Danish and Florida isolates of MdSGHV was similar with three virus delivery protocols, as a liquid food bait (in sucrose, milk, or blood), sprayed onto the flies in a Potter spray tower, or by immersiion in a crude homogenate of infected house flies. The most effective delivery system was immersion in a homogenate of ten infected flies/ml of water, resulting in 56.2% and 49.6% infection of the house flies challenged with the Danish and Florida strains, respectively. (Geden, C. J., T. Steenberg, V.-U. Lietze, and D. G. Boucias. 2011. Salivary gland hypertrophy virus of house flies in Denmark: Prevalence, host range, and comparison with a Florida isolate. J. Vector Ecol. 36: 231- 238.)

The effect of Musca domestica salivary gland hypertrophy virus (MdSGHV) on selected fitness parameters of stable flies, Stomoxys calcitrans (L.), was examined in the laboratory. Virus-injected stable flies of both genders suffered substantially higher mortality than control flies. By day 9, female mortality was 59.3 +10.1% in the virus group compared with 23.7 + 3.7% in the controls; mortality in virus-injected males was 78.1 + 3.1% compared with 33.3 + 9.3% for controls. Fecundity of control flies on days 6-9 was 49-54 eggs deposited per live female per day (total, 8,996 eggs deposited), whereas virus-injected flies produced four to five eggs per female on days 6-7 and less than one egg per female per day thereafter (total, 251 eggs). Fecal spot deposition by virus-injected flies was comparable to controls initially but decreased to +50% of control levels by day 4 after injection; infected flies produced only 26% as many fecal spots as healthy flies on days 6 and 7. None of the virus-injected stable flies developed Page 79 symptoms of salivary gland hypertrophy. Quantitative real-time polymerase chain reaction demonstrated virus replication in injected stable flies, with increasing titers of virus genome copies from one to four days after injection. MdSGHV in stable flies displayed tissue tropism similar to that observed in house fly hosts, with higher viral copy numbers in fat body and salivary glands compared with ovaries. Virus titers were 2 orders of magnitude higher in house fly than in stable fly hosts, and this difference was probably due to the absence of salivary gland hypertrophy in the latter species. Geden, C. J., A. G. Maruniak, V.-U. Lietze, J. Maruniak, and D. G. Boucias. 2011. Impact of house fly salivary gland hypertrophy virus (MdSGHV) on a heterologous host, stable fly (Stomoxys calcitrans). J. Med. Entomol. 48: 1128-1135

See attached pdfs for complete texts of the above papers .

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FINAL RESEARCH CO-ORDINATION MEETING On “Improving SIT for Tsetse Flies through Research on their Symbionts and Pathogen”

Vienna, Austria 26-30 March 2012

AUTHOR (S): Drion Boucias, Vereena Lietze. Christopher Geden, and Zhang Hu.

TITLE OF WORKING PAPER: Studies on the Hytrosavirus infecting the house fly Musca

domestica.

SHORT SUMMARY OF THE RESEARCH

Our studies on the Musca domestica initially focused on the detection, extraction, and characterization of the causal agent of salivary gland hypertrophy. This research and studies on the pathology, transmission, and epizootiology have been summarized in prior reports and published in a series of manuscripts. Significantly our research has provided a platform for the comparative analysis to the GPSGHV leading to the eventually creation and naming of the virus family Hyrosaviridae and the two viral genera Muscavirus and Glossinavirus. The second line of research conducted during the five-year period focused on the abundance and detection of this virus in Musca domestica and other related fly species. Analysis of flies collected from around the world demonstrated that this virus has a global distribution and to date is found in only Musca domestica populations. In most populations the incidence of the virus is relatively low 0-40%. However, once the virus once is established at a site it is capable of maintaining itself in the fly population over long time periods. For example, dairies from which the virus was isolated initially in 1992 was found to contain viremic house flies in the 2008 sampling.

Present evidence suggests that this virus, released in salivary secretions during feeding, can be transmitted orally to healthy conspecifics. It is assumed that ingested virus particles enter the insect via the midgut and are transported to the salivary gland. The efficiency of oral

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transmission is not high and likely is hindered by the well-developed adult peritrophic matrix that serves as a barrier for viral ingress. The primary site of viral replication is the salivary gland, the major tissue supporting morphogenesis. Recently, using qPCR methods, reported DNA and transcripts in non-salivary gland tissues. However, electron microscopy of these tissues demonstrated that the virus was limited to tracheal cells that permeate all insect tissues, and no evidence of viral replication was observed in these tissues. Unfortunately, attempts failed to replicate MdHV in available insect cell lines (unpublished data). One of the unique features of MdHV infection is that it causes rapid cellular and nuclear hypertrophy in the salivary glands within 2-3 days post-infection. This virus, unlike the well-studied insect baculovirus group, lacks a biphasic life cycle and infects non-dividing, differentiated adult cells. Viremic gland cells do not undergo apoptosis but remain intact, continuously producing and shedding enveloped virus into the gland lumen over the adult lifespan. Electron micrographs of infected cells demonstrated that the nucleocapsids are produced in the nucleus, exit the nucleus, become enveloped within the cytoplasm, migrate to, and bud out of the plasmalemma bordering the gland lumen.

The published research conducted over the last 18 months focused on the following topics:

1. Tissue tropism of the Musca domestica salivary gland hypertrophy virus. The tissue tropism of Musca domestica salivary gland hypertrophy virus (MdSGHV) infecting adult house flies was examined by transmission electron microscopy (TEM) and quantitative real- time PCR. TEM demonstrated that characteristic MdSGHV-induced nuclear and cellular hypertrophy was restricted to the salivary glands. Both nucleocapsids and enveloped virions were present in salivary gland cells. In contrast, thin sections of midguts, ovaries, abdominal fat body, crops, air sacs and brains showed the presence of enveloped virions in vacuoles of tracheal cells associated with these tissues. However, no sites of viral morphogenesis were detected in the tracheal cells. Quantitative analysis of MdSGHV DNA and transcript titers revealed that viral DNA was present in all hemolymph and tissue samples collected from MdSGHV-infected flies. Average numbers of MdSGHV genome copies per 50 ng of DNA varied significantly between examined tissues and ranged from 3.83 x 10 8 (± 3.75 x 10 7) in salivary gland samples to 7.98 x 10 5 (± 2.91 x 10 5) in hemolymph samples. High levels of viral genome copies were detected in midgut, fat body and brain samples. Viral transcripts were present in all examined samples, and transcript abundance was also at the highest level in salivary glands and at the lowest level in hemolymph. However, over the range of different Page 83

tissues that were analyzed, there was no correlation between estimated quantities of genome copies and viral transcripts. The function of viral transcripts in host tissues that do not show sites of viral morphogenesis remains to be elucidated. 2. Disease dynamics and persistence of MdSGHV-infections in laboratory house fly (Musca domestica ) populations. Past surveys of feral house fly populations have shown that Musca domestica salivary gland hypertrophy virus (MdSGHV) has a world-wide distribution with an average prevalence varying between 0.5% and 10%. How this adult-specific virus persists in nature is unknown. In the present study, experiments were conducted to examine short-term transmission efficiency and long-term persistence of symptomatic MdSGHV- infections in confined house fly populations. Disease rates transmitted from virus-infected to healthy flies in small populations of 50 or 100 flies ranged from an average 3% to 24% and did not vary between three tested isolates that originated from different continents. Introduction of an initial proportion of 40% infected flies into fly populations did not result in epizootics. Instead, long-term observations demonstrated that MdSGHV-infection levels declined over time resulting in a 10% infection rate after passing through ten filial generations. In all experiments, induced disease rates were significantly higher in male flies than in female flies and might be explained by male-specific behaviors that increased contact with viremic flies and/or virus-contaminated surfaces. 3. The cytopathologies of the two hytrosaviruses, MdSGHV and GpSGHV, in their respective host insects. Recently, a new virus family (Hytrosaviridae) was proposed for double-stranded DNA viruses that cause salivary gland hypertrophy in their dipteran hosts. The two type species, MdSGHV and GpSGHV, induce similar gross pathology and share several morphological, biological, and molecular characteristics. This histological study revealed profound differences in the cytopathology of these viruses supporting their previously proposed placement in different genera. 4. Analysis of the structural proteins from the Musca domestica Hytrosavirus with an emphasis on the major envelope protein. Studies were directed at identifying key structural components of the viral envelope and nucleocapsid. SDS-PAGE of detergent- treated virus fractions identified protein bands unique to the envelope and nucleocapsid components. Using prior LC-MSMS data we identified the viral ORF associated with the major envelope band, cloned and expressed recombinant viral antigens, and prepared a series of polyclonal sera. Western blots confirmed that antibodies recognized the target viral antigen

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and provided evidence that the viral protein MdHV96 underwent post-translational processing; antibodies bound to the target high molecular weight parent molecule as well as distinct sets of smaller bands. Immunogold electron microscopy demonstrated that the anti- MdHV96 sera recognized target antigens associated with the envelope. The nucleocapsids migrated from the virogenic stroma in the nucleus through the nuclear membrane into the cytoplasm, where they acquired an initial envelope that contained MdHV96. This major envelope protein, appeared to incorporate into intracellular membranes of both the caniculi and rough endoplasmic reticulum membranes and mediate binding to the nucleocapsids. Oral infection bioassays demonstrated that the anti-HV96 polyclonal sera acted as neutralizing agents in suppressing the levels of orally acquired infections.

In addition to the published data, research on the Musca domestica hytrosavirus (MdHV) includes studies that involves the analysis of the viral and salivary gland transcriptome, disruption of peritrophic membrane with reducing agents, analysis of the brush border membrane proteins, and the characterization of the ORF86 and its involvement in nucleocapsid trafficking and envelope formation.

In summary our research on Muscavirus over the past five years has laid the foundation for the development of this virus as an model system of a virus that infects fully differentiated adult cells leading to the shut down of metabolic pathways required for egg production in insects. Secondly the approach and technologies from our studies have been used as aframework for studying and comparing this virus to its related Glossinavirus .

______RETURN THE COMPLETED FORM TO: Ms. Magali Evrard, Insect and Pest Control Section, International Atomic Energy Agency, Room A-2155, P.O. Box 100, A-1400 Vienna Austria. Fax: +43 1 26007,e-mail: [email protected] NOT LATER THAN 29 February 2012 . (Preferably in electronic form). ______Please bring with you to the meeting your complete working paper together a copy of the abstract and working paper on diskette in Word .

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Final report on the IAEA Research Contract N° 14320

1. Contractor: Professor Flobert Njiokou University of Yaounde I, BP 825, Yaounde , Cameroon e-mail : [email protected] Tel: 237 77 71 96 31 2. Research team : Flobert Njiokou, Jean Arthur Mbida Mbida (Lectureur), Farikou Oumarou (PhD student), Kame Ngasse Ginette Irna (Master student) Gérard Cuny and Anne Geiger

3. Title of the project: Tripartite interaction between the tsetse fly, its endosymbiont and trypanosomes: an investigation on field populations of flies sampled in foci of the sleeping sickness in Cameroon

4. Objective : To study the relation between the presence of the symbiont Sodalis glossinidius or some genotypes of this bacteria and the presence of trypanosomes in tsetse flies. Page 86

5. Work planning: A - Detection of trypanosomiasis foci, and entomological prospection B – Identification of trypanosome species on Glossina species C – Investigation on Sodalis glossinidus D – Verification of the existence of a correlation between vectorial competence of Glossina and Sodalis glossinidus

6. Five year results: A- Sleeping sickness detection and entomological prospection - The foci considered were Bipindi and Campo in 2008, Fontem in 2009 and Bipindi again in 2011. - Sleeping sickness detection, in connection with the National control program of HAT (Table 1)

Foci Number of subject Number of Prevalence of Sleeping analysed positive sickness (%) Bipindi 2008 3783 3 0.08 Campo 2008 3528 6 0.17 Fontem 2009 8409 4 0.05 Bipindi 2011 4652 1 0.02 Total 20372 14 0.07

- Entomological prospection Prospection was done in many villages of the following foci, Bipindi and Campo in 2008, Fontem in 2009 and Bipindi again in 2011. Glossina palpalis palpalis was the only tsetse fly species found in Bipindi and Fontem while in Campo, 94% of flies were G. p. palpalis and 6% grouped G. nigrofusca, G. pallicera and G. caliginea . The density of flies, the teneral and the infection rates are given in Table 2. These parameters vary among the foci, villages and seasons.

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Table 2: Density and Infection rate of tsetse flies Locality Number of Density % of teneral Midgut Proboscis flies flies infection rate infection rate (%) (%) Bipindi 596 1.27 4.19 1.76 8.92 (2008) Campo 1743 4.81 16.12 1.27 5.68 (2008) Fontem 812 2.94 15.6 2.20 - (2009) Bipindi 351 3.25 12 0.3 - (2011)

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B – C: Prevalence of Sodalis glossinidus and trypanosome species All the tsetse fly samples analysed, harboured the symbiont Sodalis glossinidus , the prevalence varying from 45% to 69% (Table 3). Parasites belonging to T. brucei s.l. complex, T. b. gambiense subspecies and T. congolense species were also found. The infection rates varied among the foci, the villages and the seasons.

Table 3. Prevalence of Sodalis glossinidus and trypanosome species The number of positive PCR is given with the prevalence in the brackets; Prev. : prevalence

Locality Number Prev. Prev. T. Prev. T. b. Prev. T. Prev. T. of G. p. Sodalis brucei s.l. gambiense congolense congolense palpalis glossinidus “forest type” “savannah analysed (%) type” Bipindi 225 145 (64.4) 45 (20) 13 (5.7) 60 (26.6) 36 (16) 2008 Campo 225 102 (45.3) 20 (8.8) 2 (0.9) 34 (15.1) 29 (12.9) 2008 Fontem 157 101 (64.33) 34 (21.66) 15 (9.6) 47 (30) 24 (15.28) 2009 Bipindi 225 156 (69.33) 17 (7.6) 0 (0) 14 (6.2) 27 (12) 2011 Total 832 504 (60.57) 116 (13,9) 30 (3.6) 155 (18.62) 116 (13.9)

D – Level of association of Sodalis glossinidus infections and trypanosome infections In general, Table 3 shows strong association between the presence of the symbiont Sodalis glossinidius and the presence of the parasites. These associations were highly significant in Bipindi 2008, Campo 2008 and Fontem 2009 but not in Bipindi 2011.

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Table 3. Level of symbionts/parasites association - The null hypothesis is no association between S. glossinidus and parasites - An association occurs when the odds ratio value is superior to 1 - There is a significant association when the p value is inferior to 0.05

Locality Level of association among Sodalis glosinidus and different parasites T. brucei s.l. T. brucei T. congolense T congolense gambiense forest type savannah type Bipindi - odds ratio - odds ratio - odds ratio: - odds ratio: 2008 : 3.7 : 16.4 3.2 1.8 - p < 0.001 - p < 0.0001 - p < 0.001 - p = 0.14 Campo - odds ratio: - odds ratio: - odds ratio: - odds ratio: 2008 61.3 6.14 2.19 4.55 - p< 0.0001 - p = 0.056 - p = 0.039 - p < 0.0001 Fontem - odds ratio: - odds ratio: - odds ratio: - odds ratio: 2009 0.481 0.113 0.376 1.098 - p< 0.087 - p< 0.006 - p< 0.012 - p = 0.839 Bipindi - odds ratio: - - odds ratio: - odds ratio: 2011 2.25 1.70 5.80 - p = 0.667 - p = 0.312 - p = 0.152

7. Comment of results: - Cameroon belongs to the hypo-endemic area of HAT in the Africa, what is confirmed by the low prevalence of HAT in Bipindi, Fontem and Campo during the five year of stydy. - Only Glossina palpalis palpalis was captured in Bipindi and Fontem while four species occur in Campo, also dominated by G. p. palpalis . - The density of flies and their midgut infection varied among the foci, the villages and the seasons. - In Bipindi, the parasite infection rates have significantly decreased from 2008 to 2011, probably due to HAT control activities. - The statistical tests performed show that there is a significant association between S. glossinidus and almost all the trypanosome species tested. These results are interpreted as the existence of a positive action of the symbiont on the trypanosome installation in the midgut. The lack of significant association between S. glossinidius and the parasites in 2011 may be due to the very low prevalence of these parasites, for example 0% T. b. gambiense .

8. Conclusion During the five year study, three HAT foci were investigated. In every focus, entomological prospections were conducted in three villages in two seasons. The tsetse fly Page 90

species were identified. The infection rates of Sodalis and trypanosome species were studied using molecular techniques. Associations between Sodalis and trypanosome species were performed. The workplan was then fully followed.

9. Perspective Investigation on Glossina fuscipes fuscipes , HAT vector in the Eastern Region of Cameroon.

10. Publication relative to the subject

- Farikou O., Njiokou F ., Mbida Mbida J.A., Njitchouang G.R., Nana Djeunga H.C., Tazoacha A., Simarro P.P., Cuny G. & Geiger A. 2010. Tripartite interactions between tsetse flies, Sodalis glossinidus and trypanosomes – An epidemiological approach in two historical human African trypanosomiasis foci in Cameroon. Infection, Genetics and Evolution . 10: 115-121.

- Farikou O., Njiokou F ., Simo G., Tazoacha A., Cuny G. & Geiger A. Tsetse fly blood meal modification and trypanosome identification in two sleeping sickness foci in the forest of southern Cameroon. 2010. Acta Tropica , 116: 81-88.

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WORKING PAPER –CRP2012

Investigations on the presence and effects of Wolbachia and the salivary gland virus (SGHV) in laboratory and wild populations of Glossina pallidipes and G. austeni

Florence Wamwiri, Samuel Guya and Robert Changasi

Kenya Agricultural Research Insitute – Trypanosomiasis Research Centre (KARI-TRC) PO Box 362-00902 Kikuyu

Abstract

We report the finding of the salivary gland virus that causes hypertrophy and reproductive abnormalities in G.austeni , G.pallidipes and G.brevipalpis from nine populations in Kenya. A total of 773 flies comprising 414 G.pallidipes , 269 G.austeni and 90 G.brevipalpis were analysed. Infection prevalence obtained were 41.9%, 72.9% and 18.6% respectively. Laboratory colony SGHV analysis revealed about 13% infection in G.pallidipes . Wolbachia infection in G.pallidipes was nil whereas it was fixed (100%) in G.austeni . We further studied the co-infection of trypanosomes with the virus in both natural and experimentally-infected populations. Natural infection was evaluated from sympatric G.pallidipes and G.austeni collected from the coastal belt. Experimental infection was assessed by in vivo infection of teneral G.pallidipes males using the isolates TBR KETRI3738, TBB KETRI3386 and TC KETRI3805. Results indicate virus infection rates of 18.4% and 36.4% for G.pallidipes (N=141) and G.austeni (N=182) respectively, while trypanosome infections rates were 16.3% and 19.2% respectively. The proportion of trypanosome infection was higher in virus-negative than in virus-positive flies, however this was not significant. In the experimentally-infected flies, infection rates of 70.3%, 10% and 10% respectively were established. Experimental flies were SGHV-negative despite the colony flies having a global infection of about 13%. These preliminary results point to virus infection having a negative correlation with trypanosome infection. Future studies will focus further on infection studies and will attempt experimental clearing and re-infection with virus innoculum.

Introduction

Glossina pallidipes is one of the most important species of tsetse flies in Africa. In Kenya, it is of great economic importance as the most widely distributed species and as a vector of animal trypanosomosis. SIT is one effective method of controlling T&T but success of this method is constrained by presence of the salivary hyperplasia gland in rearing facilities. The prevalence of this virus in the field is unclear as most of the reported work was based on morphological features. The availability of new primers will enable us carry out PCR confirmation of these prevalence.

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Tsetse control can also be achieved by manipulation of naturally-occurring symbionts such as Wolbachia which may influence the reproductive capacity of the fly through the CI phenomenon. . CI has been demonstrated in Glossina morsitans (Alam et al , 2011). However, previous studies with wild G.pallidipes from Kenya and Uganda detected no Wolbachia infections (Cheng et al. , 2000). It is necessary to investigate Wolbachia presence and strains in order to investigate whether CI is expressed in other species.

The ultimate goal of this CRP was to contribute to the improvement of the sterile insect technique (SIT) for tsetse flies through research on their symbionts and pathogens. The overall objective was to understand and exploit interactions between tsetse flies and their microbes so as to enhance the efficacy and implementation of tsetse programs with an SIT component. The specific objectives of our participation in this CRP were:

1. To investigate the occurrence of the SGHV in Kenyan G.pallidipes populations 2. To determine the prevalence and strains of Wolbachia in both laboratory and field populations 3. To demonstrate the extent of CI-mediated sterility in field and laboratory crosses

Main findings (a) SGHV infection prevalence: TRC G.pallidipes colony showed a prevalence of about 13%, but no hypertrophy was detected by dissection. Field dissections also yielded about 1.5% (N= 210) hypertrophied glands, and this was only in one population Nguruman. Overall, infection was found all the tsetse belts studied except in the western Kenya area Busia. Virus infection was highest in Shimba Hills G.austeni flies. Detailed results are presented in Table 1 below:

G.pallidipes G.austeni G. brevipalpis Location N n% N n% N n% Busia 30 0 - - - - Kiboko 20 15 - - - - Meru National Park 50 0 - - - - Mwea Reserve 10 0 - - - - Nguruman 111 13.5 - - - - Arabuko-Sokoke - - 84 29.7 59 18.6

Shimba Hills N.R 193 13.4 185 43.2 31 0

Grand Total 414 41.9 269 72.9 90 18.6

(b) Wolbachia infection and typing

Wolbachia infection in G.austeni was fixed (100%) in 182 samples analyzed, whereas it was 0% in 100 G.pallidipes samples analyzed from both field and laboratory colonies. Wolbachia typing

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revealed identity between Wolbachia strains from the two coastal sites identical to GenBank wAus (Glossina austeni ) AF020077. The identity held with wsp typing of the four hypervariable regions (HVRs) of the wsp gene sequence. The HVR peptides obtained in this study were equivalent to peptide numbers 80, 40, 210 and 18 (allele 507). MLST typing revealed a new sequence type 197.

(c) Trypanosome and virus infection in G.pallidipes and G.austeni at the Kenya coast

Prevalence (%)

Species Source N V+ T+ V+T+ V-T+

G.pallidipes Shimba 141 18.4 16.3 5.7 9.9

G.austeni Shimba 132 40.9 21.2 5.3 15.9

Sokoke 50 24 14.0 6.0 8.0

182 36.3 19.2 5.5 13.7

Table 2: Trypanosome (T) and virus (V) infection in G.pallidipes and G.austeni from Shimba Hills and Sokoke. Prevalence values shown as % of total analyzed

Other activities carried out within the Project

i. Prevalence of the secondary symbiont Sodalis was also assessed in all the samples that were analyzed. Field infections of Sodalis were very low (<10%) whereas laboratory collections had high infections (>70%) which can be attributed to the membrane feeding protocol used in the colonies.

Departures from level of activity foreseen

We were unable to proceed to our objective 4 i.e. to demonstrate the extent of CI-mediated sterility in field and laboratory crosses due to the following: - Our finding of no Wolbachia infections in neither the G. pallidipes lab nor field populations - Unavailability of G.austeni from TTRI to provide pupae for establishment of the laboratory colony during the course of the CRP

(b) Publications resulting

1. Vangelis et al., (2012) Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina ), BMC Microbiology , 12 (Suppl 1):S3 doi:10.1186/1471-2180-12-S1-S3

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2.

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Testes flies dynamics, distribution and biology in west of Africa Ouédraogo / Sanon Gisèle Marie Sophie, Anicet Georges Ouédraogo, Issa Sidibé, Jean Baptiste Rayaissé, Adly M.M. Abd-Alla,

Laboratoire National d’Elevage, 01 BP 7026 Ouagadougou 01, Burkina Faso.1

Université Polytechnique de Bobo Dioulasso

CIRDES, 01 BP 454 Bobo-Dioulasso, Burkina Faso.

FAO-IAEA Agriculture and Biotechnology Laboratory entomology unit

A b s t r a c t

Trypanosomiase transmitted by tsetse flies continue to constitute a major limiting rural development in vast area of West Africa. 3 groups of tsetse occur in West Africa (palpalis, palpalis palpalis and morsitans).

Many tsetse control campaign projects were carried out with positive impact. (Hendrickx et al. , 2004; Bauer et al. , 1999; Cuisance et al. , 1984). But these successes were not sustainable, due to lack of follow up. Currently, a PATTEC (Pan African Tsetse and Trypanosomosis Eradication Campaign) project is ongoing and may lead to the use of the sterile insect technique (SIT).

Tsetse flies are known to harbor three distinct bacteria endosymbiont; In Glossina pallidipes, the Salivary Gland Hypertrophy Virus (SGHV) is responsible for a typical teratogenic symptom of salivary gland hyperplasia (SGH) affecting both males and females and leading to infertility in the affected insects (Jura et al., 1988 ; Sang et al.,1996, 1997 and 1988). The virus has been implicated in the reduction of colony.

Indeed work was conducted in Burkina Faso on wild flies caught in various provinces of the country. In total 1433 flies from four species ( Glossina palpalis gambiensis, Glossina tachinoides, Glossina morsitans submorsitans and Glossina medicorum) were caught, of which 296 were dissected and 544 used for screening. Out of 296 flies dissected, all species inclusive, 23 flies had symptomatic GpSGHV, corresponding to 7.8%. Glossina palpalis gambiensis had the highest percentage of infection of 14. 9%. Twenty-two (22%) of 544 flies (118 flies), all species inclusive, were infected when PCR was performed. The prevalence of infetion was highest (37%) in G. palpalis gambiensis. Among the localities, Houet was the most infected with a prevalence of 59.4%.When the gene P74 Nes 2F was used, the analysis of samples with the Seibersdorf samples show that one sequence from Glossina tachinoides had two SNPs G91/A and A337/G. Page 96

KEYWORRDS

Tsetse species, West Africa, Trypanosome, Biologie, SIT, symbionts, pathogen, SGHV

Contents

1-Introduction on tsetse in West Africa

2- The major tsetse species in West Africa

3-Trypanosome problem in human and animal in West Africa

4- Method of tsetse control in West Africa Including the attempt of using SIT

5- Tsetse symbiont and pathogen in West African tsetse species including SGHV

6- Prevalence and distribution of SGH in tsetse species in West Africa

7- conclusion

8-references

1. INTRODUCTION

African trypanosomes constitute an important set of parasites that affect both humans and animals. They are caused by trypanosomes, transmitted mainly by tsetse flies (Diptera: Glossinidae). In Humans, the disease progress inexorably to death if not treated. According to the World Health Organization (WHO)(2006), The number of cases was estimated at between 50 and 70 miles. In 2009, the number of new cases as dropped below the 10 000 patients (Simarro et al., 2011). If sleeping sickness is one of neglected diseases in nowadays, the African Animal Trypanomiasis affecting livestock is a major obstacle to the development of Agriculture in West Africa. About 60 million of ruminants are at risk (FAO, 2002).

The control of the disease includes the use of drugs against the parasite and control of the vector. Control of the vector using traps and trap/target-impregnated insecticides is very effective but its use by local communities has sometimes failed. Use of drugs to control trypanosomiasis is expensive and often associated with serious side effects and resistance. In addition, the country has to spend considerable amount of foreign currencies each year to import drugs. For example in 2001, the import of drugs to control trypanosomiasis increased to nearly 500 million FCFA (762,195 Euros), which represented 60% of the total imports of veterinary drugs (internal communication, DGSV, 2001)., Page 97

Many tsetse flies from the 3 groups of tsetse occur in West Africa. However, as described by Cuisance and de la Rocque, 1980, only few have medical and veterinary interest. From the palpalis group, we can note G. tachinoides , G.palaplis gambiensis and G. palpalis palpalis occurring in degraded forest, gallery forest and mangrove while from the morsitans group we can note G. morsitans submorsitans and G. longipalpis .

Since the Eighties there have been many projects aimed at controlling trypanosomiasis and campaigns were carried out in the country but little success has been achieved (Hendrickx et al. , 2004; Bauer et al. , 1999; Cuisance et al. , 1984). The International Livestock Research and Development Center in sub humid region (CIRDES) using the SIT technique has managed to eradicate the tsetse fly in Sidéradougou agro-pastoral region in Burkina Faso.

Despite the potential success of tsetse SIT programs, several constraints need to be addressed so as to enhance efficiency and expand areas where sterile males can be used. The interaction of the tsetse fly with its symbionts and pathogens represent some of these constraints and are targeted in the present proposed. Pathogens of tsetse interact with SIT in two ways, one negative and one positive. Firstly the presence of a virus can compromise the productivity of laboratory colonies and so interferes with effective mass rearing, and secondly fungal pathogens may be used as a means to suppress tsetse populations prior to the release of sterile males.

The observation of abnormal mortalities due to virus infection is well documented in mass rearing of medically and/or economically important arthropods. In Glossina pallidipes , the Salivary Gland Hypertrophy Virus (SGHV) is responsible for a typical teratogenic symptom of salivary gland hyperplasia (SGH) affecting both males and females and leading to infertility in the affected insects (Jura et al., 1988 ; Sang et al.,1996 ; Sang et al., 1997 ; Sang et al., 1998). The virus has been implicated in the reduction of colony. The virus replicates in the salivary gland, the female milk gland, midgut ovaries and testes and is known to be transmitted both vertically (transovum transmission) and sexually (veneral transmission). Virus specific primers are now available for this virus (Abd-Alla Adly et al.,2007).

Three species of tsetse flies, namely G. m. submorsitans, gambiensis G. palpalis, G. tachinoïdes are still being reared at CIRDES laboratory since the SIT eradication campaign in Sidéradougou agro-pastoral region. The presence of tumor buds form was observed on the ventral side of the abdomen of both male and female flies from the colony recently the same observation was made in glossina palpalis gambiensis from Seibersdorf. The symptoms were first observed on wild G. m. submorsitans in Burkina Faso. Page 98

The present research aims a better understanding of the relationship between flies and pathogen in the view of their use against tsetse in a sterile male release technique program.

2. The major tsetse species in West Africa In type Glossina you have three groups or sub-types: Austenina or group fusca (13 species and 4 sub-species); Nemorhina or group palpalis (5 species and 7 sub-species); Glossina or group morsitans (5 species and 3 sub-species). Morphologically glossinas are recognizable by the cell médico-slipped disc in the form of ax at the level of wings. Difference of wings size was determined between Glossina palpalis Gambiensis of Senegal (small size) and those of the Burkina Faso. (Solano and al, on 1999) this difference can be explains by weak pluviometer in Senegal. In Guinea investigation showed in 2005 presence of glossina palpalis gambiensis. A morphological study and genetic showed that there is a difference between the same populations of glossinas coming from tree different places in Guinea.

3-Trypanosome problem in human and animal in West Africa

One of the main challenges which West Africa is still facing in that third millennium is to achieve an adequate level of food security for a growing population while maintaining sustainable natural resources. An important contribution to solving that challenge is expected from the livestock sector, one of the pillars of the economy. Trypanosomiasis problem constitutes there a major concern for the producers and important quantities of money are used each year to face the problem.

In Burkina Faso for example the livestock sector contributes to sustained food security and nutrition, and plays for nearly 12% of GDP and is around 24% of total exports by value, not taking into account its contribution to the animal traction, transport and land organic manure. The national herd is 7.5 million head of cattle. The growth rate is estimated at 2% for cattle (ENECII, 2004). Trypanosomiasis is particularly important in the west part of the country, where more than 60% of livestock are under the risk to contract disease (Kruska et al. , 1995).

African trypanosomes are transmitted to mammals cyclically by tsetse flies. However, some species of trypanosomes such as T. evansi, T. vivax (Hoare, 1972; Desquesnes & Dia, 2003b) and T.congolense (Desquesnes & Dia, 2003a) can be Page 99

transmitted mechanically by other insects such as tabanids (Atylotus agrestis ), the T Stomoxys etc. .. . equiperdum is the only species sexually transmitted. The degree of animals’ infection with trypanosomes depends on the trypanosome species involved but also to the vertebrate host (Table 2). The cycle of T. vivax development is about 14 days, but variable going from 7 to 40 days (Dale et al ., 1995). T. brucei spp . is transmitted after a long cycle approximately 30 days, but it can vary between 17 - 45 days (Hoare, 1970)

In general, all tsetses can cyclically transmit trypanosomes. However, small groups of 12 to 15 species or sub – species, due to the fact of their distribution are major vectors of HAT and AAT. In addition to the species listed above, tsetse of morsitans group ( G. morsitans submorsitans and G. longipalpis ) are also vectors of AAT in West Africa and transmit Trypanosoma vivax , T. congolense and T. brucei brucei .

West Africa is composed by 14 countries: Burkina Faso, Benin, Mali, Ivory Cost, Gambia, Ghana, Guinea, Guinea Bissau, Liberia, Mali, Niger, Nigeria, Senegal, Sera Leone and Togo. Tsetse of Palpalis group ( Glossina tachinoides , G. palpalis gambiensis and G. palpalis palpalis ), are the essential vector of HAT, transmitting Trypanosoma brucei gambiense ( Jamonneau, 2004 )

The results of medical prospection made between 2000 and 2006 with National programme of country and financial support of OMS, France and Belgium cooperation and IRD.

The figure 2 shows that in Burkina Faso, Benin, Mali and Togo no cases were reported despite active and passive surveillance (World Health Organization, 2008; Simarro PP. et al., 2008).

Suspected cases reported in Benin between 2000 and 2003 and from Mali between 2000 and 2002 were not confirmed.

Cases reported in Burkina Faso between 2000 and 2004 have been labeled and attributed to Ivory Cost the Country of infection (Cecchi G., et al).

Data for Guinea include revisions of previously published figures for the years from 2002 to 2004 (World Health Organization, 2008; Simarro PP. et al., 2008).

For Gambia, Senegal Sere Leone, Niger, Liberia and Guinea Bissau no data was reported.

4. Method of tsetse control in West Africa including the attempt of using SIT

The control of the disease includes the use of drugs against the parasite and control of the vector. Control of the vector using traps and trap/target-impregnated insecticides Page 100

is very effective but its use by local communities has sometimes failed. Use of drugs to control trypanosomiasis is expensive and for example in 2001, the import of drugs to control trypanosomiasis increased to nearly 500 million FCFA (762,195 Euros), which represented 60% of the total imports of veterinary drugs (internal communication, DGSV, 2001). In addition, drugs are often associated with serious side effects and nowadays, resistance cases are frequent (Geerts & Holmes, 1998; Simbarashe et al ., 2011). In nowadays, la lutte intégrée demeure l’option de choix (Vreysen, 2001).

In the fit against the binomial tsetse-trypanosomes, knowledge on their correlations remains very important. To understand complex correlations between tsetse flies and trypanosomes a geographical frame, is essential on epidemiological plan and constitutes a key element for the rational and efficient installation of the measurements of control of trypanosomoses (Aksoy, 2003).

Studies on correlations trypanosomes-glossines-pathogène introduce considerable challenges owing to difficulties in l 'élevage and the assertion of the flies in big number and because of the weakness of the rates of ripening of the parasite and of necessary time to supplement cycle in the fly.

Many tsetse control campaign projects were carried out in Burkina Faso since the years eighties with positive impact. (Hendrickx et al. , 2004; Bauer et al. , 1999; Cuisance et al. , 1984). Unfortunately, these successes were not sustainable, due to lack of follow up. Currently, a PATTEC (Pan African Tsetse and Trypanosomosis Eradication Campaign) project is ongoing and may lead to the use of the sterile insect technique (SIT).

In Senegal International Agency for Atomic Energy projected to eliminate tsetse flies in Niayes. In Mali the project will covers 37,000 km2, aims to reduce the population of tsetse flies about 19,000 km2 in Baní and eliminate about 18,000 km2 in Niger. It will increase significantly the production of milk, meat and cereals.

5- Tsetse symbiont and pathogen in West African including SGHV

However, despite the potential success of tsetse SIT programs, several constraints need to be addressed so as to enhance efficiency and expand areas where sterile males can be used. The interaction of the tsetse fly with its symbionts and pathogens represent some of these constraints. Pathogens of tsetse interact with SIT in two ways, one negative and one positive. Firstly the presence of a virus can compromise the productivity of laboratory colonies and so interferes with effective mass rearing (Ellis & Maudlin, 1987; Jura et al ., 1989), and secondly fungal pathogens may be used as a means to suppress tsetse populations prior to the release of sterile males (Rio et al. , 2004). Page 101

The main endo symbiont Wigglesworthia of the tsetse fly are in the cells of the intestine and Sodalis glossinidius in salivary glands and midgut. Several studies have shown the importance of these symbionts in the physiology of the fly. Thus it was proved by Dale & Welburn (2001) that the elimination of S. glossinidius increases the refracting nature of the tsetse fly against trypanosome infection (T b rhodesiense). In addition, the discovery of specific genotypes of S. glossinidius may be related to vector competence of tsetse flies (Geiger et al., 2007) argue for a role of the symbiont in the tsetse-trypanosome interactions but the mechanisms underlying these interactions are said to understand.

The observation of abnormal mortalities due to virus infection is well documented in mass rearing of medically and/or economically important arthropods. In Glossina pallidipes, the Salivary Gland Hypertrophy Virus (SGHV) is responsible for a typical teratogenic symptom of salivary gland hyperplasia (SGH) affecting both males and females and leading to infertility in the affected insects (Jura et al., 1988 ; Sang et al.,1996, 1997 and 1988). The virus has been implicated in the reduction of colony. The virus replicates in the salivary gland, the female milk gland, midget ovaries and testes and is known to be transmitted both vertically and sexually. Specific primers are available for this virus (Abd-Alla Adly et al., 2007).

Through monitoring of Glossina breeding in CIRDES, tumors buds form of serving on the ventral side of the abdomen of the flies were observed under a binocular microscope (photo 3). These remarks were made first on Glossina morsitans submorsitans females and females of wild Glossina palpalis gambiensis . Several cases of salivary gland hypertrophy were observed in tsetse flies from the Sissili province in Burkina Faso.

The present research is aimed at a better understanding of the interrelationship between flies and pathogen in the view of their use in programs against tsetse based on sterile males release technique.

6- Prevalence and distribution of SGH in tsetse species in West Africa (case of Burkina Faso)

6.1 Material and Methods

Six (6) provinces of high tsetse and trypanosomosis presence (Banwa, Comoé, Houet, Kénédougou, in the western part and Sissili and Tapoa respectively in the Southern and Eastern parts of the country) were covered by the study.

Flies were collected using Biconical Challier-Laveissière and monoconical Vavoua traps deployed within 200-m radius in 24 hours. They were sorted by species and each subsample of 50 flies was transferred into 15-ml tube containing 95% alcohol. Each tube was labeled with a pencil on a piece of paper inserted inside and another one was glued outside. The label included location, date, specie, and GPS geo-references. Page 102

After 24 hours, alcohol was poured and replaced with fresh and clean one. The tube was sealed with a cap and parafilm and repacked.

Samples were brought to the laboratory where flies were dissected ( Yoni et al., 2006) to determine the prevalence of salivary glands hypertrophy.

In order to determine the prevalence of GpSGHV virus by PCR samples was sensed in FAO-IAEA Agriculture and Biotechnology Laboratory entomology unit , the DNA was extracted from caught flies using qiagen protocol .

The PCR reaction was performed using in a final volume of 25 µl, using 2 master mix buffer Primers Mix2 (82R+83F; GpCAG 133R+F; GpSHV 2R+F) 1µl 0, 5 µl, Total DNA 1, 5 µl, in order to check the presence of the virus in our samples, the presence of genomic DNA and to know if there is DNA in our samples.

PCR machine was under the following reaction conditions; 35 cycles and Annealing at 58°C (Abd-Alla, et al., 2007). The PCR product was detected with Agarose (1%) gel electrophoresis and ethidium bromide staining.

Up to 1433 flies were caught in total, from which 329 were dissected and 544 used for screening We performed nested PCR with 6 genes (P74 primers, ODV-e66 primers, Pif-1 primers, Pif-2 primers, Pif-3 primers, DNA pool primers), cleaned the PCR products and sent for sequencing for gene verification.

For the study of papillae on the ventral side of the abdomen of tsetse flies we used four protocols that have been established on the species Glossina morsitans submorsitans taken from the breeding colony of CIRDESS. To determine frequencies of flies with papillae according to age class, we isolated 100 flies and observed them every 10 days under a binocular microscope.

Some essays for experimental contamination were tried. For this some lesions were dissected, crushed in physiological water and put into contact with other flies.

The crushed liquid was also used to make culture using gelose in order to isolate bacteria.

6.2. Results

For the realization of this work we were particularly interested in the Western and the southwest part of the Burkina Faso where Trypanosomose is particularly important. We crossed all in 8 provinces it is about the province of Houet, about Komoé ( Folenzo), about Tapoa (Diapaga, Singou, Pama), of Kossi ( solenzo ), Kénédougou and Sissili ( léo ). Page 103

The proportion of flies captured was as follows: 32.40, 29.15, 23.90 and 14.53 % for Glossina papalis gambiensis (Gpg), Glossina tachinoides (Gt), Glossina morsitans submorsitans (Gms) and Glossina medicorum (Gmed), respectively. Glossina morsitans submorsitans and G. medicorum were captured only in Folonzo, in the Comoé province (figure 3).

6.2.2. The proportion of flies captured by locality

The figure 4 shows us the proportions of flies captured according to the locality. 48,29 % of the captured flies result from the province of Komoé, 12,39 % of Kossi, 12 % in Houet, 11,51 % in Tapoa, 7,78 in Sissili and 7,38 % in Kénédougou.

6.2.3. The proportion of flies infected by Gp SGHV

From the 296 flies dissected, all species comprised, 23 flies were found with symptomatic Gp SGHV, giving a global prevalence of 7,77 %. Glossina palpalis gambiensis was the species with the highest percentage of infection with 14,87%, the prevalence was 6,66 and 0% respectively for G. m. morsitans and G. tachinoides (figure 4). Dissection was not made of G. medicorum .

Up to 118 flies all species included out of 544 were infected when we performed PCR, hence a global infection rate of 22.34%. When considering the species individually, the prevalence is more than 10% for each of them, and is even up to 37% for G. palpalis gambiensis (figure 4)

When considering the localities, the Houet and Kenedougou ones were the most infected with a prevalence of 59 and 56% respectively, up to 59.42%, while no case was detected in the Sissili, and only 3.57 and 4.44% in the Tapoa and Banwa provinces respectively (Figure 5).

The analysis of samples with ODV –e66 primers Bac 1F, DNA POL Nes-1F, PIF1 Nes- 1F, PIF3 Nes-1F did not show any mutation (n=10, n=8, n=7and n=7 respectively).

With the gene P74 Nes 2F; we don’t get the right size. But we sensed the samples for gene verification. The Analysis of samples from Glossina morsitans submorsitans, Glossina tachinoides, Gossina medicorum , and Glossina palpalis gambiensis Seibersdorf samples show that one sequence from Glossina tachinoides have two SNPs G91/A and A337/G. this sequence id 44-7C529 (figure 6).

Analysis the samples from BKF from Gms, Gt, G.med , with the Seib samples did not show any mutation (n=3) with PIF 2 Nes-2F but the sequence of 94-2D533 should be repeated several times from both ends.

6.2.4. Papilla lesions and mortality in breeding flies by age class Page 104

Detection of abdominal lesions using microscopic examination indicated an increase of number of flies with lesions with age class of breeding flies (figure 7). The first cases of flies with lesions are observed at 30 days. Increase of cases numbers seems to be due to a contamination process.

Some cases of mortality have been observed during the study but they may be due to manipulation of flies (Figure 7). It could not be concluded that there is an association between observed lesions and mortality cases.

6.2.5. Essays of transmission of lesions and isolation of pathogenic agent

The essays of transmission from flies to flies using crushed lesions solution didn’t give any positive result. However, numerous bacterial colonies were obtained from culture using gelose substrate but that result was not useful because it was not possible to distinguish bacterial colonies. Further studies should used gradients of diluted crushed lesions for bacterial culture. Specific culture substrates for bacteria and fongic organics should be used also for better study of papilla etiology. Samples of solution for experimental contamination should also be taken from flies that weren’t fed during two days to avoid interrelations between host pathogenic agents and those of the fly.

The percent of flies with papilla lesions in glossina palpalis gambiensis held in Seiberdorf is very important 55% of old flies observed during my training was positives. Former studies indicated that lesions were found in all Glossina species met in Burkina Faso with an exception for Glossina medicorum . For Glossina palpalis gambiensis and Glossina morsitans submorsitans the papilla lesions are dark patches and/or white papillas which are abundant. These lesions differ from coupling scars (See photo 2, 3, 4, 5). For Glossina tachinoïdes, mainly one patch is observed. The lesion affected both males and females. There was not case with simultaneous abdominal lesion and salivary gland hypertrophy.

7-conclusion

Considering the prevalence of 0.5-5% generally observed in wild population, Otieno et al., 1980, one could say that the proportion of up to 22.34% is high. Considering the fact that males presenting an hypertrophy of the salivary glands are completely sterile, and that females have a decline of the fertility (Jura et al., 1988; Sang et al.,1996,1997 and 1998), the productivity of the colony of breeding for the technique of release sterile males can be reduced.

It would be interesting to envisage a biological fight by taking into account this virus especially that this pathology can affect quite the species of glossines (Gouteux, J.P., 1987; Odindo et al.,1982). We know that this virus can be passed on horizontally and vertically (Jura et al .,1989), the males contaminated could be Page 105

envisaged with the aim of reducing the productivity of the wild glossines prior to the release of sterile males.

For papilla lesion observed, experiments can be realized on Glossina palpalis gambiensis Colonie from Seibersdorf.

Our preliminary results suggest that transmission of the pathogenic agent may not be by way of contact. The ingestion way may be investigated: crushed lesions solution may be injected in the abdomen of flies or administrate through the flies meal.

The investigations may include analysis of more factors for example;

-Coupled flies should be used to determine numbers of pupas per day along the time in comparison to those of healthy flies;

- Some dissections of dead females’ flies to determine the reproductive status (uterus content, abortion, presence of follicular reliquary, ovary status) should be done;

-The numbers of cases by sex group should be determined.

One can say that this pathology have to be taking in count in PATTEC program because often pupas are involved from CIRDESS to Start a new breeding.

The extension of fly’s collection to new areas to provide more representatively of flies’ samples from the other country from West Africa will be use full. In addition, new factors like flies sex, season of capture, areas, morphology and genetic variability between the same population of flies will be taken into account in the experimental design and statistical analysis.

We highly recommend the continuation of the research to achieve the objective of contribution to better understanding of interrelations between flies and their pathogen and use of these results to improve strategies against tsetse flies.

Methods and techniques used during field and laboratory activities allowed us to get the interesting preliminary results. This also justifies the necessity to follow up the present research program. The following activities are planned: 1. Collect tsetse flies from different species and population in the field in West Africa. 2. Evaluate the prevalence of SGHV in the collected flies by PCR. 3. Analyze the genetic variability of SGHV in the infected tsetse population collected from the field. 4. Write a report and scientific articles on the results achieved.

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Simarro P. P., Diarra A., Ruiz Postigo J. A., Franco J. R. & Jannin J. G. 2011. The human African trypanosomiasis control and surveillance programme of the World Health Organization 2000-2009: the way forward. PLoS Negl Trop Dis 5: 2, e1007.

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

Solano P, Kaba D, Ravel S, Dyer NA, Sall B, et al. (2010) Population Genetics as a Tool to Select Tsetse Control Strategies: Suppression or Eradication of Glossina palpalis gambiensis in the Niayes of Senegal. PLoS Negl Trop Dis 4(5): e692. doi:10.1371/journal.pntd.0000692

Van Den Abbeele J., Claes Y., van B. D., Le R. 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 ( Pt 5): 469-478.

Vickerman K. 1985. Developmental cycles and biology of pathogenic trypanosomes. Br Med Bull 41 : 2, 105-114.

Vreysen M. J. 2001 . Principles of area-wide integrated tsetse fly control using the sterile insect technique. Med Trop (Mars ) 61 : 4-5, 397-411. Page 110

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IMPROVING SIT FOR TSETSE FLIES THROUGH RESEARCH ON THEIR SYMBIONTS AND PATHOGENS

Progress report on project: “Prevalence of salivary gland hypertrophy virus (SGHV) in different tsetse populations and the possibility of establishing SGHV free tsetse colonies” CONTRACT No. 14264 / R4 (September 2011 – March 2012)

Imna Malele Tsetse & Trypanosomiasis Research Institute, P. O. Box 1026 TANGA – TANZANIA Email: [email protected] AND [email protected]

Summary The sterile Insect technique is an important component in area wide integrated tsetse control in tsetse infested areas. However presence of hypertrophied salivary glands virus in the wild tsetse which are the seeds for colony adaptations in the lab; are a stumbling block in adapting fly colonies in the lab. The virus is transmitted both vertically (in the wild) and horizontally (in the lab). Their prevalence is magnified in the lab as a result of the existing feeding regimen. Virus infection rates in wild collected flies and analysed both by microscopic and molecular technique from tsetse species of economic importance from the coastal and inland areas of Tanzania, namely Glossina fuscipes fuscipes, G. pallidipes, G. morsitans and G. swynnertoni is reported and discussed. This is based on the observation made following four years of closely monitoring the prevalence of SGHV; which shows that the virus is prevalent in various tsetse infested areas and all tsetse species named above were found infected irrespective of their ages, sex, and season of the year; although infection levels differed among species and from one location to the other. Microscopic infection was 1.2% (25/2164) from the coast as compared to 0.4% (6/1725) for inland collected flies. Molecular analysis increased the detection of infected numbers to 19.81% (104/525). Many of the flies which were found positive by molecular analysis were asymptomatic. From this observation, we conclude that when planning to initiate tsetse eradication against G. f. fuscipes, G. pallidipes , G. morsitans and G. swynnertoni other options other than SIT may be explored due to the fact that various wild populations of these species are infected by the salivary gland hyperplasia virus, and if used as seeds for initiation of the colonies, the virus could stagnate tsetse rearing and hence delay the implementation of tsetse control for the targeted species. Wolbachia are known to infect a wide range of arthropods where it causes a variety of reproductive abnormalities, one of which is cytoplasmic incompatibility that when expressed causes embryonic death due to disruption in fertilization events. Occurrence of Wolbachia was common in wild G.m.morsitans with both single and double insertions. For Sodalis , the prevalence ranged from 21.1% to 73.7% depending on the type of primers used in detections. Although their effect on TTRI lab flies have not been quantified but the two can be linked to the unsatisfactory performance of some colonies especially the G. pallidipes.

Introduction and rationale Tsetse flies are the vectors of trypanosomes, the causative organisms of trypanosomiasis: nagana, in animals and sleeping sickness in man. The disease causes death of countless domestic animals especially cattle, and therefore depriving tsetse infested areas of meat, milk and the use of animals for draught purposes (Jordan, 1986). Farmers are forced to keep their animals alive by the use of drugs mainly Samorin and Berenil. However, irregular availability of trypanocidal drugs and frequent under dosing of animals have resulted into subsequent development of trypanosome resistance to the commonly used drugs ( Geerts &

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Holmes, 1998 ). Tsetse infestation therefore renders most of fertile areas unexploited thus limiting their contribution to the economy of affected communities. Further, overgrazing and degradation of land in tsetse free areas is common as most of the cattle are confined to these areas (Malele et al ., 2011).

The human trypanosomiasis is another setback which results from the presence of tsetse in tsetse infested area. In Africa although the number of cases have dropped from 300,000 new cases reported in 1998 (WHO, 1998) to 10,000 cases; still over 60 million people living in some 250 locations are at risk of contracting the disease (WHO, 2010; Simarro et al., 2011). In addition, the presence of tsetse causes human beings to abandon their settlements. Such depopulation and cessation of farming activities normally leads to regrowth of bushes, consequently extending suitable habitats for Glossina and thus trypanosomiasis (Reid et al ., 2000). In Tanzania, tsetse species that affect both human and livestock, include, Glossina morstans sl, G.pallidipes, G. longipennis, G. brevipalpis, G. austeni, G. swynertonni and G.fuscipes sl. The common and widely distributed species throughout the country are G. pallidipes and G. m.morsitans . G. swynnertoni and G. longipennis are restricted to the north western part along the Serengeti ecosystem . G. swynnertoni is incriminated to be behind several outbreaks of trypanosomiasis of human form specifically among tourists (Jelinek et al ., 2002; Malele et al., 2007). The population of G. longipennis in the ecosystem is very low. The G. fuscipes is restricted to the riverine along lakes Victoria and Tanganyika. Whereas G. f. fuscipes is found around Lake Victoria, G. f. martini is found along the drainage systems of Lake Tanganyika. The G. brevipalpis is found in forested areas in the country whereas the G. austeni is somehow found in the coastal belt . Hence the four species; G. pallidipes, G. morsitans sl., G. swynnertoni and G. fuscipes are species of major economical importance due to their coverage (widely distributed) and their role in the epidemiology of African Trypanosomiasis.

For many years, efforts to control tsetse flies have been conducted in the country. Unfortunately, the techniques used to control tsetse have limitations and failed to provide a complete solution to the problem. These include clearing of vegetations which serve as habitats for different tsetse species (Jordan, 1986); control by chemicals in the form of aerial (Kgori et al. 2006) and or ground spraying; traps and insecticide treated targets and also by baiting animals (Fox et al. , 1991). In some areas Traps and targets have been used in combination with live baits to speed up suppression of the vector to eliminate the flies. Treatment of live baits especially cattle have involved dipping, pour-ons, spot-on, or spray- on using appropriate formulations.

The Sterile Insect Technique (SIT) is a genetic or autocidal method, which employs a pest species for its own destruction. The technique has been found extremely efficient at low populations of a target species (Knipling, 1964). The technique is based on the principle that laboratory mass-produced males are sterilized with small doses of gamma radiation and regularly released in large numbers in the field to copulate with wild female flies which, consequently, fail to produce offspring to maintain the reared colony. SIT requires the production of large numbers of flies from a rearing facility to obtain sufficient surplus males for sterilization and release in the field. Sustained and systematic releases of sterile male insects to outnumber the indigenous wild fertile males reduce the reproductive capacity of female flies of the target population resulting into its collapse. The technology works best when the target population in an area is low, hence requires other technologies to reduce the wild populations prior to the application of SIT. In Tanzania the technique has been used

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against G. m. morsitans and achieved a reduction of about 83% of the species at Mkwaja ranch in Tanga region (Williamson et al ., 1983) (1972 – 1979). The second application was on G. austeni in 1994 – 1997 in Unguja, Zanzibar Island (Msangi et al ., 2000) and the island is free from cyclically transmitted trypanosomiasis.

The need to rear tsetse in the laboratories for surplus males which are later gamma sterilized for release while females are left in the colony for breeding is the main stumbling block with SIT application. Confinement of flies in captivity makes them susceptible to diseases attributed by pathogens and symbionts (Abd-Alla et al., 2011). Although in nature they are infected by both pathogens and symbionts, the magnitude of infection is increased in the laboratory as a result of confinement in rearing cages, and feeding system. Fly rearing is by in vitro feeding (Bauer and Wetzel, 1976), where blood (fresh defibrinated bovine blood) is supplied to the flies through an artificial membrane, after warming it up to vertebrate body temperature and tsetse must pierce a membranous surface to suck blood. The blood that is supplied to the flies nourishes both sexes to be productive and the offspring in the adenotrophic viviparity of the female tsetse fly. The female tsetse produces an offspring every 9 – 10 days at a 25°C environment (Feldmann, 1994; Gooding et al ., 1997). Research has shown that membrane feeding is one of the most efficient ways of transmitting pathogens in tsetse (Abd- Alla et al ., 2011). Salivary gland hyperplasia virus (SGHV) is one of the pathogens that affects tsetse in the wild, but confinement in the lab amplifies the magnitude of virus infection (Abd-Alla et al ., 2011).

At the TTRI insectaries, both G. pallidipes and G. brevipalpis are reared. Other species included G. m. morsitans, G. m. centralis and G. austeni . Occasionally the two colonies have performed sub-optimally and there was a suspicion that infection with the SGHV might contribute to this. To answer this, it was decided to investigate and monitor the presence of SGHV in colony flies. Currently no data on the prevalence of this or related viruses in lab tsetse have been documented in Tanzania. Hence the study was conducted to determine whether our tsetse colonies are also infected by the virus.

Prevalence of SGHV in different tsetse populations in Tanzania For the past four years, research was conducted on tsetse species of G. pallidipes, G. m. morsitans, G. fuscipes fuscipes and G. swynnertoni in Tanzania to evaluate the extent of SGHV prevalence. The above mentioned tsetse species are not only species of economic importance to Tanzania but also in other tsetse-infested belts of Africa. The aim of the study was to (i) determine the prevalence of the virus infection in tsetse wild population, (ii) identify SGHV free tsetse populations that would provide fly seeds for the establishment of SGHV free tsetse colonies so as to speed up implementations of SIT programs. The anticipation is that SGHV free populations would be more productive to meet sufficient fly numbers for SIT that could not be achieved in time in SGHV infected fly populations. To determine the prevalence of the virus, both microscopic and molecular methods (Abd-Alla et al., 2007) were used.

Symptomatic infection rate of salivary gland hypertrophy (SGH) in wild tsetse A total of 3886 flies were dissected, 2164 from the coast and 1725 from inland. The infection in G. pallidipes from the coast was 1.2% (25/2164) and none for other species (163) from the same area (i.e. G. morsitans, G. brevipalpis and G. austeni ). From the inland flies, no infection was recorded in 922 G. pallidipes dissected ; 0.6% (4/706) was recorded for G. swynnertoni and 2.06% (2/97) for G. f. fuscipes (Table 1).

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Flies of all ages were found infected by the virus when examined under microscope (Fig. 1). However the number of SGH positive flies was very low hence this data might not depict the actual situation in the wild. Females which were infected by the virus had the ovulation category of 1, 2, 3, 4, 5 and 6. For males, their ages were estimated by wing fray method and it ranged from 8 to 30 days with more falling around ages of 17 – 19 days; hence the virus affected flies of different age groups. Analysis by molecular technique indicated that many flies (both sexes), were asymptomatically infected and irrespective of the season of the year.

Molecular detection of SGHV in wild tsetse flies PCR screening of SGHV infection in the wild tsetse are presented in Table 2. Different tsetse species showed different levels of SGHV infection. The infection rate in G. f. fuscipes was 18.22% (43/236); 18.75% (30/160) in G. pallidipes ; 15% (12/80) in G. swynnertoni and 55% (11/20) in G.m.morsitans. The PCR product sizes of about 400bp were detected for positive samples for GpSGHV usingGpSGHV1F and GpSGHV1R primers previously described by Abd-Alla et al., (2007) (Figure 2). The level of infection both microscopically and by molecular analysis indicated that some species had high prevalence of the virus than other species. G. pallidipes was more infected followed by G. m. morsitans and none in other species ( G. austeni and G. brevipalpis ).Flies from the coast area especially G. pallidipes appeared to be more infected than the same species from inland. The infection in G. pallidipes from Serengeti was 12.35% compared to 25% from the coast. The infection in a sympatric species ( G. swynnertoni ) was 15% (12/80). The infection in G. f. fuscipes was found to be 18.22% (43/236) although microscopically the level of infection in the species was very low (Table 2).

Documented information on SGHV which were based on microscopic observations and information obtained from this study, both shows that many species in the wild are infected by the SGHV although the levels differ amongst species, populations and from one location to the other. We have recorded different levels of SGHV prevalence from various areas in Tanzania. The overall infection rate in G . pallidipes was 18.75% (30/160) from both sites and 55% (11/20) in G. m. morsitans from the coast. Infection rates in G. pallidipes from inland were 12.35% (10/80) and in the sympatric species, G. swynnertoni, the infection was 15% (12/80). The prevalence in G. f. fuscipes was 18.22% (43/236). We recorded more infections of the virus in male G. f. fuscipes 16.95% (40/236) than in females 1.3% (3/236). All positive G.m.morsitans were male flies 55% (11/20). The exception was with the G. pallidipes from Selous where the positive samples were all female flies 25% (20/80). The infection was found in both male and female tsetse though in different infection levels. Flies which were infected had estimated ages of from 8 to 45 days old. No virus infection was recorded from G. brevipalpis and G. austeni both microscopically and by molecular analysis. However this cannot rule out the presence of the virus in the species as it has been reported already from G. austeni collected from Amani in Tanzania (Leietze et al , 2011). The reason the infection was missed here could be due to the lower numbers of tsetse that were trapped. Through molecular analysis, G. m. morsitans was found infected though it was asymptomatic. G. m. morsitans has been reared before in the insectaries but there is no information that the species had the virus, however the feeding regimen ( in vivo ) was different from what is being practised now ( in vitro ) (Williamson et al ., 1983).

Treatment against the SGHV in the laboratory includes treating the diet meant for tsetse with an antiviral compound in order to decontaminate it (Cannavan et al ., 2010). However, this might seem promising if it won’t add the cost to the mass rearing activities as already SIT is seen as one of the technique with unbearable initial costs such that many scholars tend to

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advocate other conventional techniques for control than SIT (Molyneux, 2001; Roger and Randolph, 2002; Torr et al ., 2005). Treatment of tsetse diets with antiviral drug had no negative impact on the productivity or mortality of producing females but also no significant reduction in virus copy number was accrued as a result of the antiviral drug application. Two compounds were tested: the Valacyclovir and Acyclovir. Results showed that there was a slight increase in mortality with Valacyclovir drug despite the fact that productivity was better than the Acyclovir drug (FAO/IAEA, 2009). The practicality of this compound remains pessimistic.

SGHV and its implication to tsetse control Many wild tsetse, were found infected by SGHV when analysed by molecular analysis even though they were asymptomatic (no hypertrophied glands). The virus is present in wild tsetse although many flies were asymptomatic. In wild flies the virus is not species, sex, age and season dependent but levels of infection differs from one species to the other. High infection was detected in G. f. fuscipes from inland; and in G. pallidipes and G. m. morsitans from the coast. Virus infection was high in G. pallidipes collected from the coast and relatively low in G. pallidipes from the inland site. The results suggests that when planning to initiate tsetse eradication campaigns against G. f. fuscipes and G. pallidipes , care should be taken when planning to initiate tsetse laboratory colony for SIT eradication program against the species by selecting non infected flies to be sued to initiate the colonies to minimize the infection rate with SGHV could stagnate tsetse rearing and hence delay the implementation of tsetse control for the targeted species and management of SGHV infection in the established colony should be applied.

Research on other tsetse symbionts and their relation with virus from the two study sites and lab flies were also carried out. The study showed the presence of various levels of Wolbachia and Sodalis in lab and wild flies. Wolbachia are known to infect a wide range of arthropods where it causes a variety of reproductive abnormalities, one of which is cytoplasmic incompatibility that when expressed causes embryonic death due to disruption in fertilization events (Aksoy, 2003). Some Wolbachia strain have also been found to reduce host life span for both young and older adults host insects (Weiss and Aksoy, 2011). In the lab, symbionts like Sodalis have shown to have detrimental fitness effects on infected tsetse than their wild counterparts as measured by longevity and fecundity (Weiss et al., 2006).

Wolbachia & Sodalis The occurrence of Wolbachia in the species analysed was very common. We used three types f primers Wsp, 16S RNA and FbPA as indicated in Table 4. The primers detected the Wolbachia from the three species although levels differed from one species to the other. Products by Wsp primers ranged between 500 – 632 bp; whereas for Wspec the product size was 438 bp. Sodalis infection was common in both lab and wild flies, with band sizes ranging from 400, 500 and 600bp. However high prevalence of Sodalis infection was found in G. m. morsitans compared to G. pallidipes , a sympatric species (Table 5).

Interaction with Trypanosome infection Trypanosome types obtained from wild flies were T. vivax and T. congolense type and only two flies were found infected out of 38 flies analysed from the coastal site and non from 40 G. f. fuscipes. The data obtained this year was not sufficient to make a conclusive association between trypanosomes, SGHV, and symbionts.

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Conclusions and recommendations SGHV is an issue in tsetse rearing. It affects fecundity/reproduction and survival in tsetse (Abd-Alla et al 2010). SIT has stagnated because of SGHV in tsetse colonies e.g. the Arba Minch strain of Ethiopia (Abd-Alla et al 2010). A decade has gone by since the successful control of tsetse by SIT in Zanzibar and the attempt to have another successful eradication program in the mainland have many challenges i.e. the susceptibility of the target species to the virus infection. Although SIT is a very important component in area wide integrated tsetse control, tsetse rearing in the presence of SGHV means SIT programs need the development and establishment of virus management strategy to reduce or eliminate the harmful impact of the virus on tsetse colonies of susceptible target species. In wild flies the virus is not species, sex, age and season dependent but levels of prevalence differs from one species to the other.

So far we have not been able to find any positive case from our four years study from the lab flies (Table 3); however if it happens to find a positive case then it will important to Sequence the Virus from the lab colony in order to see the similarity between viruses that have been reported in other lab fly colonies. We have monitored the virus from species of different ages, and the results recorded were nil.

Both lab and wild flies are infected with other symbionts. It will be important to gather more data on their co-infection of SGHV, Wolbachia and, Sodalis in lab flies and SGHV, Wolbachia, Sodalis and trypanosomes co- infections in the two tsetse populations from the two study sites. The two symbionts (Wolbachia and Sodalis ) occurs in both wild and lab flies, they are not species specific, neither sex dependent nor fly age. Those infected were found both during the wet and dry season. It has been proved in the lab that Wolbachia induces cytoplasmic incompatibility and this can be exploited for Insect Pest population control (Zabalou et al ., 2004).

Acknowledgements The collaborations with Prof Anna Malacrida, Prof Kostas Bourtzis, and Prof Jan Den Abbeele are very much appreciated. Mr. Kitwika, Mr. H. Nyingilili, E. Lyaruu, A. Mnyamisi for their helping hand in lab and field work. This study was funded by IAEA through IAEA/CRP project NO: 14264 and Project ID No.: A80132 of WHO/TDR. I wish to acknowledge the TTRI Director for logistic support; and Adly (IAEA) for useful guidance related to this work.

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Figure legend

Figure 1: Proportional of symptomatically infected flies per estimated fly ages in days

Figure 2: Screening of salivary gland hyperplasia virus infection using PCR. Bands at ~400bp indicate the positive samples. Numbers 1- 11 Glossina swynertoni Females, 12-20 Glossina swynertoni Males, 21- 30 Glossina pallidipes Female and 31- 40 Glossina pallidipes Male. M -

DNA ladder FastRuler; VT – positive control ex Tanzania; VV – positive control ex Seibersdorf and VC- negative control Page 120

Table 1: Prevalence of SGHV in different tsetse species by microscopic observation ZONE SITE SPECIES SEX Total + ve % Estimated ages by wing No. SGHV fray for males and ovarian category or females Coastal Msubugwe G. pallidipes F 412 4 0.97 18 M 728 5 0.69 22 Gendagenda G. pallidipes F 300 12 4.0 8, 18,45, M 461 4 0.87 19 Rufiji/ G. pallidipes F 100 0 0 Selous M 0 0 0 G. m. morsitans F 10 0 0 M 125 0 0 G. brevipalpis F 10 0 0 M 12 0 0 G. austeni F 6 0 0 M 0 0 0 Total for coast 2164 25 1.2

Inland Tarangire G. pallidipes F 268 0 0 M 193 0 0 G. swynnertoni F 208 1 0.5 8 M 127 1 0.8 17 Serengeti G. pallidipes F 300 0 0 8,19 M 161 0 0 17 G. swynnertoni F 371 2 0.54 20 M 0 0 0 Rorya G. f. fuscipes F 39 2 5.13 8, 18,45, M 58 0 0 Total for inland 1725 6 0.35 Overall total 3889 31 0.8

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Table 2: Prevalence of SGHV from tsetse species by molecular analysis

No Tsetse species District Village Asympto-matic Females Males +ve females +ve males Total +ve analysed flies

1 G.f.fuscipes Rorya Bubombi 45 0 45 0 8 8 2 Rorya Masonga 40 19 21 0 0 0 3 Ukerewe Ukerewe 40 12 28 0 2 2 4 Kagera Kemondo 40 22 18 0 0 0 Rorya Ras Nyab 71 23 48 3 38 41 Subtotal 236 76 168 3 40 43 (18.22%) G. pallidipes Rufiji Selous 80 40 40 20 0 20 (25%) Serengeti Death Valley 80 40 40 4 6 10 (12.35%)

Subtotal 160 80 80 24 6 30 (18.75%) G. swynnertoni Serengeti Death Valley 80 40 40 6 6 12 (15%)

G.m.morsitans Rufiji Selous 20 0 20 0 11 11 (55%) G.austeni Rufiji Muyuyu 15 10 5 0 0 0 G.brevibalpalis Rufiji Muyuyu 14 10 4 0 0 0 Grand Total 525 216 309 33 (15.28%) 71 104 (22.98%) (19.81%)

Table 3: PATHOGEN detection in colonized tsetse species at TTRI: Salivary Gland Hyperplasia Virus SPECIE SEX Total AGE GpSGHV RESULT FROM DIFFERENT PRIMERS number (days) P74- p74- Pif1- Pif1- Odv-E66 Bac 2F+2R per age 1F+1R 2F+2R 1F+1R 2F+2R F+R G.m.morsitans M 5 >21 ------G.pallidipes M 3 >70 ------G.pallidipes M 1 >55 ------G.pallidipes F 1 >55 ------G.austeni M 2 >98 ------G.austeni F 3 >98 ------G.m.centralis F 5 >98 ------G.brevipalips M 2 >98 ------G.brevipalips M 1 >70 ------Total 23 0/23 0/23 0/23 0/23 0/23 0/23

Table 4: Wolbachia infections in wild tsetse populations Tsetse Orig Sex Zone Molecular FbPA WSpec Wsp (F1/R1) species in TRYPS

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Gff Wild F Inland 0/40 5/40 1/40 0/40 Ras Nyabera (RNB) Gp Wild F Inland ND 11/18 0/18 (sb) 2/18 (sb) (Death (db 3/18, Sb 8/18) Valley) Gmm Wild F Coastal ND 13/18 (sb 7/18; db 13/18 db 13/18 (db 2/18; sb (Selous) 6/18) 11/18) ND: Not Done

Table 5: Sodalis detection in Wild tsetse from the coastal area Tsetse Origin Sex Zone Location Sod GroEl Sod FTZ species Gmm Wild F Coastal Selous 16/18 (88.9%) 0/18 Gp Wild F Coastal Selous 12/20 (60%) 8/20 (40%) Overall Total 28/38 (73.7%) 8/38 (21.1%)

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Working paper March 2012 Jan Van Den Abbeele

Institute of Tropical Medicine Antwerp Unit Veterinary Protozoology

Expression and extracellular release of a functional anti-trypanosome Nanobody ® in Sodalis glossinidius , a bacterial symbiont of the tsetse fly Linda De Vooght, Guy Caljon, Jan Van Den Abbeele Published in Microbial Cell Factories 2012, 11(1):23

Introduction Sodalis glossinidius is a maternally inherited gram-negative bacterial endosymbiont of the tsetse fly that can be found both inter- and intracellularly in the tsetse fly midgut, muscle, fat body, milk glands, and salivary glands [1]. Given the close proximity of S. glossinidius to the different insect tissues where trypanosome parasites reside and the fact that it is one of the few insect symbiotic bacteria that can be cultured and genetically modified in vitro [1-3], S. glossinidius is considered as a potential in vivo drug delivery vehicle to control T. congolense and T. brucei development in the fly. This strategy involving the use of bacterial symbionts to express foreign proteins, designed to block pathogen transmission, is often referred to as paratransgenesis [4] and has been developed and proposed to combat different insect–borne animal and human diseases [5-7]. The control of the trypanosome parasite in the tsetse fly using paratransgenic technology requires the identification and characterization of gene products that interfere with trypanosome development without reducing the fitness of the tsetse host or its symbiont. Nanobodies ® (Nbs) represent the smallest known intact antigen-binding fragments derived from heavy-chain only antibodies (HCAbs), devoid of light chains, naturally occurring in Camelidae and sharks [8-10] (reviewed by [11]). Due to their ability to target unique epitopes that are less well targeted by conventional antibodies, Nbs are currently of high research interest for various pharmaceutical applications, including diagnosis and drug delivery [12,13]. Because of their superior intrinsic properties, e.g. small size (13-15 kDa), strict monomeric behavior and high in vitro stability [9,14] Nbs are efficiently produced in micro-organisms such as Escherichia coli [15] and therefore show high potential as effector molecules in the paratransgenesis approach. Nbs directed towards distinct regions of the variant-specific surface glycoprotein (VSG), abundantly present on the surface of bloodstream trypanosomes, have already been identified, targeting VSG epitopes that are inaccessible on live trypanosomes for larger antibodies [16]. An essential aspect for a successful symbiont-based paratransgenesis approach is the active release of the effector molecules to the inner insect environment for efficient targeting of the pathogen. However, few studies have focused on the export of heterologous proteins to the periplasmatic and/or outer environment of S. glossinidius .

We investigated the potential of S. glossinidius to express functional Nbs without interfering with cell viability. Secondly, we evaluated the capability of two independent secretion signals, predicted to be involved in different protein secretion pathways, to deliver the effector proteins to the extracellular environment. We show that Nb_An33, Page 124

recognizing a VSG epitope on Trypanosoma brucei [16] was expressed by S. glossinidius . Furthermore, we demonstrated that the pectate lyaseB (pelB) signal peptide from Erwinia carotovora is able to direct the export of fully functional Nb_An33 to the periplasm of S. glossinidius resulting in significant levels of extracellular release. Importantly, Sodalis strains that efficiently released the effector protein were not affected in their growth, suggesting that they may be competitive with endogenous microbiota in the midgut environment of the tsetse fly.

Main Results & Discussion Genetically modified bacterial symbionts of arthropod disease vectors are potential tools for the delivery of proteins that interfere with pathogen development in the vector and may serve as a powerful complementary approach to control disease transmission [2]. Furthermore, the use of bacterial symbionts expressing foreign proteins in disease- carrying arthropods has also an intriguing potential for studying insect-pathogen interactions. The advent of Nanobody ® technology has offered new prospects for the development of new effector molecules applicable for the paratransgenesis approach. These single-domain antigen-binding fragments represent exquisite targeting tools because of their small size (13-15 kDa) and stability properties [9,14]. Despite the interest for a paratransgenesis approach in tsetse flies to control transmission of African trypanosomiasis, little progress has been made on the identification and expression of trypanosome-interfering proteins in the tsetse fly bacterial endosymbiont Sodalis glossinidius . To date, S . glossinidius strains have only been used as hosts for the production of GFP [1]. In this study we explored the possibility of expressing a trypanosome-interfering Nanobody ® in Sodalis glossindius . We have developed a suitable expression vector that allows for the expression in S. glossinidius of an anti- trypanosome Nanobody ®, Nb_An33 that targets a high-mannose carbohydrate epitope present on the Variant-specific Surface Glycoprotein of Trypanosoma brucei [16]. Importantly, to control parasites in the tsetse fly, it is imperative for the effector molecules to reach their target. In an effort to address the need for an efficient secretion system we evaluated two distinct bacterial secretion pathways in their capacity to deliver Nbs to the extracellular environment. Nb_An33 fused to the S. glossinidius FliC secretion signal was expressed by both S. glossinidius and E. coli . However, in neither species could the fusion protein be detected in the culture medium nor the periplasm. The underlying reasons for this secretion failure remains unclear. One possibility includes the presence of a functional Sodalis glossinidius flagellar cap protein FliD, responsible for the polymerization of the FliC monomers into a filament, which would hinder the release of the FliC-fusion protein into the extracellular medium [17].

Nb_An33 was successfully exported to the S. glossinidius and E. coli periplasm using the pelB signal peptide. Furthermore, in both species the recombinant protein accumulated in the culture medium with an extracellular export efficiency in Sodalis of 20-25% during exponential and stationary growth phase where cell lysis is negligible. The extracellular release of small proteins and antibody fragments that were secreted into the periplasm via the pelB signal peptide has already been described [7,18]. The occurrence of this phenomenon appears to be highly dependent on the characteristics of the protein and is not yet fully understood. The secretion of some recombinant proteins to the periplasm is suggested to cause a destabilization of the outer membrane, which becomes leaky and results in the non-specific release of periplasmic proteins to the extracellular environment [19,20]. The pelB signal peptide is known to direct Page 125

protein translocation to the periplasm via the Sec-dependent type II secretion pathway [21]. The feasibility of this pathway to export heterologous proteins to the periplasm of S. glossinidius complements the twin-arginine translocation (Tat) pathway, also involved with periplasmic transport, which was previously demonstrated to be functional in the Sodalis glossinidius [22].

Additionally, the ELISA results indicated that during the stationary and cell lysis phase of the growth curve, S. glossinidius cells harboring the ppelBNb33 lac plasmid released functional Nb_An33 in the culture medium where it accumulated at concentrations as high as 160 ng/ml. This clearly indicates that the release of Nbs from lysing Sodalis glossinidius cells to the extracellular environment could also be an important way for the expressed Nb to reach its trypanosome target in the lumen of the tsetse fly. For both S. glossinidius strains harboring pNb33lac and pFliCNb33, the amount active Nb_An33 expression was below the detection limit in ELISA, probably due to the fact that biological activity is dependent on correct protein folding involving the formation of disulfide bonds which is unlikely to occur in the reducing environment of the cytoplasm. The ELISA results also demonstrated that the secreted Nb_An33 was perfectly functional in terms of antigen binding to purified soluble AnTat 1.1 VSG in vitro . Another important consideration when expressing potential effector proteins into the midguts of blood feeding arthropods is the susceptibility of the effectors to proteolytic degradation. Spiking of midgut extracts from tsetse flies with purified Nb_An33 in an ELISA assay demonstrated that Nb_An33 retains its antigen binding properties (unpublished results), providing preliminary evidence that Nb_An33 might remain functional within the midgut environment of the fly. Furthermore, Nbs can easily be mutagenized and selected for increased proteolytic stability [23]. Finally, the capacity of the secreted Nb_An33 to recognize its epitope on living trypanosomes was confirmed by flow cytometry and fluorescence microscopy using alkaline binding conditions that are relevant for the tsetse fly midgut physiology. Growth curve analysis and cell population doubling time of the S. glossinidius strains harboring the different expression constructs showed that there was a strong correlation between the ability of the S. glossinidius strains to export the Nb_An33 fusion protein and growth performance. Strains expressing Nb_An33 intracellularly showed a significant reduction in growth rate compared to the WT strains, while secreting strains were not affected in their growth. These results suggest that accumulation of Nb_An33 in the cytoplasm imparts a detrimental effect on growth performance and that efficiently secreting Nb_An33 to the periplasm rescues this effect, allowing the strain to grow with kinetics similar to the WT strain.

Conclusions

This study provides the first demonstration of the functional expression and extracellular delivery of trypanosome-interfering proteins in S. glossinidius . Moreover, we demonstrated that S. glossinidius expressing pelBNb_An33 exhibited no significant reduction in terms of fitness, determined by in vitro growth kinetics, compared to the wild-type strain. This ability of the recombinant S. glossinidius strain to effectively compete with native strains is of great importance to the overall success of the paratransgenesis strategy. Given the ability of S. glossinidius to express high levels of active Nb_An33 and the capacity to release this anti-trypanosome Nb without hampering the bacterium viability, the foundation has been laid for further exploration Page 126

of the inhibitory effect on trypanosome development in the tsetse fly. For this, highly potent trypanolytic Nbs have been developed very recently that lyse trypanosomes both in vitro and in vivo by interfering with the parasite endocytic pathway [24].

The current study also reinforces the notion for the potential of S. glossinidius to be developed into a paratransgenic platform organism. At a broad level, the concept of using Nbs as effector molecules to be delivered by bacterial endosymbionts is not limited to the tsetse fly–trypanosome model but could also be applied in a paratransgenic approach to encompass other vector-borne diseases.

Acknowledgments

This research was supported by the Belgian Co-operation (Directorate-General for Development Co-operation, DGD), ITM SOFI-B grant, the InterUniversity Attraction Pole programme (IAP) and the ERC-Starting Grant ‘NANOSYM’. This work is also performed in the frame of a FAO/IAEA Coordinated Research Project on “Improving SIT for tsetse flies through research on their symbionts and pathogens”.

References

1. Cheng Q, Aksoy S: Tissue tropism, transmission and expression of foreign genes in vivo in midgut symbionts of tsetse flies. Insect Mol Biol 1999, 8: 125-132. 2. Beard CB, O'Neill SL, Mason P, Mandelco L, Woese CR, Tesh RB et al .: Genetic transformation and phylogeny of bacterial symbionts from tsetse. Insect Mol Biol 1993, 1: 123-131. 3. Welburn SC, Maudlin I, Ellis DS: In vitro cultivation of rickettsia-like-organisms from Glossina spp. Ann Trop Med Parasitol 1987, 81: 331-335. 4. Hooper LV, Gordon JI: Commensal host-bacterial relationships in the gut. Science 2001, 292: 1115-1118. 5. Aksoy S, Weiss BL, Attardo G: Transgenesis and the management of vector-borne diseases. In Paratransgenesis applied for control of tsetse transmitted sleeping sickness. Volume 627 . Edited by Aksoy S. Texas: Landes Bioscience; 2008:35-45. 6. Durvasula RV, Gumbs A, Panackal A, Kruglov O, Aksoy S, Merrifield RB et al .: Prevention of insect-borne disease: an approach using transgenic symbiotic bacteria. Proc Natl Acad Sci U S A 1997, 94: 3274-3278. 7. Bisi DC, Lampe DJ: Secretion of anti-Plasmodium effector proteins from a natural Pantoea agglomerans isolate by using PelB and HlyA secretion signals. Appl Environ Microbiol 2011, 77: 4669-4675. 8. Hamers-Casterman C, Atarhouch T, Muyldermans S, Robinson G, Hamers C, Songa EB et al .: Naturally occurring antibodies devoid of light chains. Nature 1993, 363: 446- 448. 9. Muyldermans S: Single domain camel antibodies: current status. J Biotechnol 2001, 74: 277-302. 10. Flajnik MF, Deschacht N, Muyldermans S: A case of convergence: why did a simple alternative to canonical antibodies arise in sharks and camels? PLoS Biol 2011, 9: e1001120. 11. Van Bockstaele F, Holz JB, Revets H: The development of nanobodies for therapeutic applications. Curr Opin Investig Drugs 2009, 10: 1212-1224. Page 127

12. Cortez-Retamozo V, Backmann N, Senter PD, Wernery U, De BP, Muyldermans S et al .: Efficient cancer therapy with a nanobody-based conjugate. Cancer Res 2004, 64: 2853-2857. 13. Saerens D, Stijlemans B, Baral TN, Nguyen Thi GT, Wernery U, Magez S et al .: Parallel selection of multiple anti-infectome Nanobodies without access to purified antigens. J Immunol Methods 2008, 329: 138-150. 14. Dumoulin M, Conrath K, Van MA, Meersman F, Heremans K, Frenken LG et al .: Single- domain antibody fragments with high conformational stability. Protein Sci 2002, 11: 500-515. 15. Arbabi GM, Desmyter A, Wyns L, Hamers R, Muyldermans S: Selection and identification of single domain antibody fragments from camel heavy-chain antibodies. FEBS Lett 1997, 414: 521-526. 16. Stijlemans B, Conrath K, Cortez-Retamozo V, Van XH, Wyns L, Senter P et al .: Efficient targeting of conserved cryptic epitopes of infectious agents by single domain antibodies. African trypanosomes as paradigm. J Biol Chem 2004, 279: 1256-1261. 17. Majander K, Anton L, Antikainen J, Lang H, Brummer M, Korhonen TK et al .: Extracellular secretion of polypeptides using a modified Escherichia coli flagellar secretion apparatus. Nat Biotechnol 2005, 23: 475-481. 18. Georgiou G, Segatori L: Preparative expression of secreted proteins in bacteria: status report and future prospects. Curr Opin Biotechnol 2005, 16: 538-545. 19. Sandkvist M, Bagdasarian M: Secretion of recombinant proteins by Gram-negative bacteria. Curr Opin Biotechnol 1996, 7: 505-511. 20. Jonasson P, Liljeqvist S, Nygren PA, Stahl S: Genetic design for facilitated production and recovery of recombinant proteins in Escherichia coli. Biotechnol Appl Biochem 2002, 35: 91-105. 21. Choi JH, Lee SY: Secretory and extracellular production of recombinant proteins using Escherichia coli. Appl Microbiol Biotechnol 2004, 64: 625-635. 22. De Vooght L, Caljon G, Coosemans M, Van den Abbeele J: Functional analysis of the twin-arginine translocation pathway in Sodalis glossinidius, a bacterial symbiont of the tsetse fly. Appl Environ Microbiol 2011, 77: 1132-1134. 23. Harmsen MM, van Solt CB, van Zijderveld-van Bemmel AM, Niewold TA, van Zijderveld FG: Selection and optimization of proteolytically stable llama single-domain antibody fragments for oral immunotherapy. Appl Microbiol Biotechnol 2006, 72: 544-551. 24. Stijlemans B, Caljon G, Natesan SK, Saerens D, Conrath K, Perez-Morga D et al .: High affinity nanobodies against the Trypanosome brucei VSG are potent trypanolytic agents that block endocytosis. PLoS Pathog 2011, 7: e1002072.

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JKI - Institute for Biological Control. Darmstadt, Germany: Jehle, J.

Collaborators: Abd-Alla, A., Bergoin, M., Boucias, D., Burand, J., Vlak, J,

The sequencing of the genomes of GpSGHV and MdSGHV provided detailed genetic information about the two viruses. The genome of GpSGHV is a double-stranded circular closed DNA molecule of 190,032 bp encoding 160 predicted open reading frames (ORFs) of more than 50 amino acids (Abd-Alla et al., 2008; 2009 ). Its A+T content of 72% is one of the highest of any insect viruses. It contains one inverted repeat sequence and 14 direct repeat sequences composed of head-to-tail tandem repeats of 51 to 246 bp. Eleven of these direct repeat sequences are clustered in two genome regions between 86.1-88.2 kbp and 179.4-183.5 kbp. The genome of MdSGHV is 124,279 bp in length with an A+T content of 46.5% (Garcia-Maruniak et al., 2008). A total of 108 ORFs were predicted for MdSGHV. The genome of MdSGHV contains 18 direct repeat sequences consisting of tandem repeats of 9 to 149 bp. The repeats are more or less equally distributed throughout the genome and less clustered as those of the GpSGHV genome.

When the genomes of GpSGHV and MdSGHV were compared to other dsDNA viruses a significant number of gene homologues with baculoviruses and nudiviruses were identified, suggesting a relatedness of these viruses. These gene homologues include twelve so-called baculovirus core genes involved in transcription (lef-8, lef-9, lef-4, lef-5), DNA replication (DNA polymerase, helicase) and the infection process (pif-1, pif-2, pif-3, p74) and others functions (ac81, p33). Strikingly, the MdSGHV encodes a homologue of a polyhedrin/granulin gene of members of the baculovirus genera Alpha-, Beta-, Gammabaculovirus (Wang and Jehle, 2009; Wang et al., 2011, 2012). Based on genome sequence comparison, degenerate oligonucleotides were designed for a rapid PCR detection of GpSGHV in tsetse flies (Abd-Alla et al., 2011).

Phylogenetic analyses of GpSGHV and MdSGHV using single and multiple gene trees resulted in conflicting results depending on the type and number of genes as well as the tree inferring methods applied (Abd-Alla et al., 2008; Wang and Jehle 2009; Wang et al. 2011; Wang et al., 2012; Jehle et al., 2012). It is proposed that SHGVs, baculoviruses and nudiviruses share a common ancestor and are related phylogenetically.

The genome sequences also supported the view that SGHVs are biologically and phylogenetically distinct enough from other dsDNA virus families and warrant the establishment of an own virus family. A new virus family Hytrosaviridae has been proposed Page 130 to the International Committee on Taxonomy of Viruses (ICTV) in 2011 (Abd-Alla et al., 2009). GpSGHV belongs to the newly established species Glossina hytrosavirus (Genus: Glossinavirus ), whereas MdSGHV belongs to the species Musca hytrosavirus (Genus: Muscavirus ). This taxonomic proposal was approved by ICTV in 2011.

Publications: JEHLE , J.A., WANG , Y. ABD ALLA , A. M. M. (2012). Evolution, Phylogeny and Taxonomy of Hytrosaviridae (Minireview, submitted to Journal of Invertebrate Pathology ) WANG , Y., BININDA -EMONDS , O. R. P., VAN OERS , M. M., VLAK , J. M., JEHLE , J. A. (2011). Nudiviruses give insights into the evolution of nuclear arthropod-specific large circular double- stranded DNA viruses. Virus Genes 42 , 444-456). WANG , Y., BININDA -EMONDS , O. R. P., JEHLE , J. A. (2012). Nudivirus Genomics and Phylogeny. In: Viral Genomes - Molecular Structure, Diversity, Gene Expression Mechanisms and Host-Virus Interactions, Maria Laura Garcia and Victor Romanowski (Ed.), ISBN: 978-953-51-0098-0, InTech, Available from: http://www.intechopen.com/articles/show/title/nudivirus-genomics-and-phylogeny ABD -ALLA , A. M. M., SALEM , T. Z., PARKER , A. G., WANG , Y., JEHLE , J. A., VREYSEN , M. J. B., BOUCIAS , D. (2011). Universal primers for rapid detection of hytrosaviruses. Journal of Virological Methods 171 , 280-283. WANG , Y., JEHLE , J. A. (2009). Nudiviruses and other large, double-stranded circular DNA viruses of invertebrates: New insights on an old topic. Journal of Invertebrate Pathology 101 , 187-193. ABD -ALLA , A. M. M., VLAK , J. M., BERGOIN , M., MARUNIAK , J. E., PARKER , A. G. BURAND , J. P., JEHLE , J. A., BOUCIAS D. G (2009). Hytrosaviridae : a proposal for classification and nomenclature of a new insect virus family. Archives of Virology 154 , 909-918. ABD -ALLA , A. M. M., COUSSERANS , F., PARKER , A., JEHLE , J. A., PARKER , N., VLAK , J. M., ROBINSON , A., BERGOIN , M. (2008). Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. Journal of Virology 82 , 4595-4611.

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RNAi Inhibition of HzNV-2 Pathology in Helicoverpa zea Moths

John P. Burand 1,2 , Justin Nguyen 1, and Woojin Kim 2

Departments of 1Microbiology & 2Plant, Soil and Insect Sciences University of Massachusetts – Amherst, Amherst, MA USA

Abstract:

RNA interference (RNAi) is a post- transcriptional, gene regulation mechanism found in virtually all plants and animals, which is thought to function in fine-tuning of protein levels in cells. Various RNAi approaches have been used successfully as a tool in the study of functional genomics in insects, for insect pest control and for the prevention of viral diseases in insects. Here we report on our use an RNAi strategy to block replication and pathology of the nudivirus HzNV-2, a sexually transmitted, insect sterilizing virus of moths. Virus infected Helicoverpa zea female moths have malformed reproductive tissues which appear as a large “Y-shaped” structure and are sterile. As with hytrosaviruses and baculoviruses, HzNV-2 codes for P-74, a virus structural protein which functions in the viral entry into cells, the first essential step in the infection the host. Interestingly, HzNV-2 also codes for a juvenile hormone esterase (JHE) gene which is thought to function by lowering the level of the transcriptional regulator JH in infected insect tissues. Double stranded RNA containing HzNV-2 JHE or P-74 sequences was found to limit the pathology in virus infected female moths

Introduction:

The insect sterilizing nudivirus HzNV-2 (a.k.a. gonad specific virus [GSV]) (1, 2) has several molecular and biological properties in common with the salivary gland hypertrophy virus of tsetse flies, Glossina pallidipes (GpSGHV). While GpSGHV infection of tsetse results in hypertrophy of the salivary gland and sterility of the infected fly (3, 4), HzNV-2 infections of the corn earworm moth, Helicoverpa zea causes gross malformation of the reproductive tissues and sterility of the infected host. In females this malformation includes hypertrophy of the oviduct tissues with proliferation of the cells that comprise these tissues. As a result of virus replication in these cells a large numbers of virus particles accumulate in the bursa which leads to the formation of a plug of virus blocking the reproductive opening of the female moth (5, 6). This plug then provides a source of virus which is transferred to healthy males that attempt to mate with infected females (7).

Since HzNV-2 and the hytrosavirus, GpSGHV have several similar biological properties, and share orthologs of several structural protein genes like p-74 (8, 9, 10), we have been using HzNV-2 as a model to study several of the biological and molecular properties of hytrosaviruses. As part of this work we have used a RNA interference (RNAi) approach to down regulate the expression of viral genes specifically the p-74 gene, to determine their role in the replication and pathology of the virus.

RNAi is a highly conserved mechanism of post-transcriptional regulation of gene expression which is believed to play an important role in fine-tuning protein levels in a wide Page 132 range of cell types (11, 12, 13). In insects there are two main classes of small non-coding RNAs (sRNAs) which are used to produce functional RNAi. These are microRNAs (miRNAs) and short interfering RNAs (siRNAs). Although miRNAs and siRNA differs in the way they enter the RNAi pathway, they both have common elements (Figure 1). Both miRNAs and siRNAs are generated from long double stranded RNAs (dsRNA) by Dicer, a ribonuclease III enzyme,

Figure 1: Pathway of RNAi biogenesis from miRNA and siRNA in the cell cytoplasm. accompanied by a transactivating response RNA-binding protein (TRBP). One strand of either type of dsRNA (referred to as the guide strand), along with TRBP is then associated with an Argonaut family protein (AGO) (14) becoming part of RISC, the RNA induced silencing complex. RISC loaded with a guide strand RNA functions to bind to specific mRNA targets determined by the guide, and inhibit translation of the message. The mRNA targets are comprised of sequences complementary or partially complementary to miRNA or siRNA sequences. Complete complementary results in the mRNA being degraded while partial complementarity results in inhibition of initiation of translation or dissociation of ribosomes. Ultimately the end result is either case is the targeted down regulation of gene expression.

RNAi has been employed in functional genomics studies in insect species from a variety of different orders, including Diptera, Coleoptera, Lepidoptera, Isoptera, Orthoptera, Hymenoptera and Hemiptera, and has proven successful in the characterization of the function of genes involved in several physiological processes in insects including: reproduction, development, behavior and immunity (15, 16).

The success of Bt crop plants has paved the way for the development of RNAi based, insect resistant, genetically modified plants (17). The gene target specificity inherent to RNAi promises to allow for the design of insect pest control strategies which are tailor-made for use against specific pests without having adverse effects on non-targets and beneficial insects (18). The species specific insecticidal effects of RNAi was demonstrated using shown dsRNAs Page 133 designed to target the vacuolar ATPase genes ( v-ATPase ) of either T. castaneum, M. sexta, A. pisum or D. melanogaster all of which resulted in significant mortality only in the conspecific species (18). Using an RNAi approach directed against the v-ATPase of the Colorado potato beetle (CPB) Baum et. al. (2007) (19) found that RNAi targeting the CPB V-ATPase was 10 times more active than dsRNA complementary to western corn rootworm (WCR), Diabrotica virgifera virgifera , v-ATPase when feed to CPB. In this same study corn plants expressing RNAi directed against the v-ATPase gene of the WCR affected RNAi activity to this gene in the insect and the transgenic plants exhibited a resistance to feeding damage by this lepidopteron pest (19).

RNAi strategies have also been used for the control of diseases in insects. Recently, field trials were conducted to demonstrate the applicability of an RNAi approach to protect bees from infection by the Israeli Acute Paralysis Virus (IAPV). In this study, IAPV specific RNAi was used as a food additive for overwintering bees in order to prevent mortality and improve the overall health of bees infected with IAPV. As a result over twice as many bees in the RNAi treated, IAPV infected hives survived to adulthood and these treated hives produced three times more honey than the untreated, virus infected hives (20).

Tsetse flies ( Glossina spp.) serve as vectors for several Trypanosome species which are the causative agents of African sleeping sickness in humans and the cattle disease Nagana in sub- Saharan Africa. Attempts at utilizing sterile insect techniques to control this insect pest have been thwarted by viral infections in colonies of laboratory-maintained flies. In wild populations of flies, the virus, GpSGHV is transmitted vertically from the mother to her offspring. In laboratory colonies where a large number of insects share a common blood meal source, the virus can be transmitted horizontally via ingestion of blood contaminated by salivary secretion from infected flies. Viral levels in laboratory reared flies can reach as high as 85%, eventually leading to the collapse of the colony. Several methods including antiviral antibodies, drugs and RNAi directed against specific viral gene targets have been used to suppress virus levels in laboratory colonies of flies with limited success (21).

Here we have used an RNAi approach to target the expression of two different HzNV-2 genes in viruses infected moths in attempt to block virus replication and pathology. These included RNAi directed against the gene that encodes the virus structural protein P-74, a protein which functions in the viral entry into cells, and the orf 7 gene which codes for a juvenile hormone esterase (JHE) gene which is thought to function by lowering the level of the transcriptional regulator JH in infected insect tissues.

Methods

Identification of putative siRNA sequences in Hz-2V, P74 and ORF7 gene sequences.

The strategy employed to inhibit HzNV-2 replication in infected insects was to block the expression of the viral p-74 or the orf 7 gene using an RNAi strategy. This was accomplished by synthesizing a ~ 700 bp long dsRNA complementary to a specific region of these genes which could be used to produce siRNAi capable of guiding the RISC to complementary, target gene mRNA sequences for cleavage and degradation ultimately leading to down regulation of the expression of these genes. Regions in HzNV-2, ORF 7 (Figure 2) and P-74 (Figure 3) genes containing putative siRNA target sequences were identified using RNAi software from Page 134 mekentosj.com ( http://mekentosj.com/irnai/ ). Next, primer sets flanking these sequences were designed for the amplification of viral DNA fragments that could sever as templates for the in vitro synthesis of dsRNAs.

Figure 2: Putative siRNA sites in the Hz-2V, ORF7 gene. The sequences hi- lighted in light grey are the best siRNA site, and those in darker grey are the least likely effective siRNA sites. Primer sets (circled) were then chosen to encompass the best putative siRNA sites.

Figure 3: Putative siRNA sites in the Hz-2V, P-74 gene. The sequences hi- lighted in light grey are the best siRNA site, and those in darker grey are the least likely effective siRNA sites. Primer sets (circled) were then chosen to encompass the best putative siRNA sites.

PCR Amplification of DNA Templates from Viral DNA

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PCR products used as templates for in vitro transcription of dsRNA were produced using the HzNV-2 genome as template and primer sets identified as outlined above, designed such that one primer in each pair contained T-7 promoter sequences (Table 1). Each reaction contained 0.1ug of viral DNA template and 30 umoles of each of the two primers. The PCR conditions were: 2 min at 94 °C followed by 40 cycles (30 s at 94 °C, 30 s at 60 °C, 45 s at 72 °C) and a final extension step at 72 °C for 7 minutes.

Table 1. Primers sets used to produce DNA templates for T7 transcription of long dsRNAs. Underlined sequence designates the T7 promoter region of one primer in the pair.

Production of Long dsRNAs

The DNA product for each of the two PCR reactions was purified using a Qiagen QIAquick PCR purification kit. Each of the two DNA products then served as a template for the synthesis of a single strand RNA using the Promega RiboMax T7 transcription system. The reactions were treated with RNase free DNase and the RNA strands combined, heated to 95 oC, and cooled to room temperature allowing them to anneal and form dsRNA molecules (Figure 3).

Denature at 95 oC & Anneal at Rm Temp

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Figure 4: Method for producing dsRNAs from two separate in vitro transcription reactions.

Infection of H. zea larvae and treatment with dsRNAs

Fifth instar H. zea larvae, were injected with 2ul of purified HzNV-2 and these infected insects were injected with long dsRNAs containing sequences complementary to the p- 74 or orf 7 gene to examine the ability of these RNAs to inhibit viral replication and/or pathology. After infection, experimental larvae were injected with ~10 ug of the dsRNA. Insects injected with 4 ul of dH 2O served as uninfected controls. Upon emergence of adult moths, females were dissected and examined for malformation of their reproductive tissues indicative of viral replication.

Results and Discussion

As expected, females injected with HzNV-2 as 5-6 instars had grossly malformed reproductive tissues as has been previously described (4). While healthy females were found to have many egg filled ovariols which appeared normal (Figure 4A), infected females had enlarged common and lateral oviducts with only a few ovariols remaining (Figure 4B). Virus infected females, treated with dsRNA (Figure 4C and D respectively) had reproductive tissues which looked very similar to those of the water injected controls. These results strongly indicate that both p -74 and orf 7 dsRNA injected into virus infected insects showed RNAi activity and inhibited the expression of their homologous HzNV-2, genes and as a result reduced viral pathology. The absence of viral pathology in HzNV-2 infected females treated p -74 dsRNA provides a proof of principle that dsRNA molecules containing GpSGHV p-74 sequences may be useful in limiting viral replication and reducing the pathology of tsetse flies infected with this virus allowing for their use in sterile release programs designed to control this insect pest.

Figure 5: Reproductive tissues from normal (A) and infected female moths (B) compared with those from infected females treated with of P74 (C) or ORF7 (D) dsRNA showing differences observed in oviducts (OD) and ovariols (OV) in these insects.

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References 1. Raina, A. K., and J.R. Adams. 1995. Gonad-specific virus of corn earworm. Nature 374:770. 2. Hamm, J.J., J.E. Carpenter and E. L. Styer. 1996. Effect of oviposition day on incidence of agonadal progeny of Helicoverpa zea infected with a virus. Annals of the ESA 89:266- 275. 3. Coler, R.R., Boucias, D.G., Frank, J.H., Maruniak, J.E., Garciacanedo, A. & Pendland, J.C. 1993, "Characterization and Description of a Virus Causing Salivary-Gland Hyperplasia in the Housefly, Musca-Domestica", Medical and veterinary entomology, vol. 7, no. 3, pp. 275-282. 4. Kokwaro, E.D., Nyindo, M. & Chimtawi, M. 1990, "Ultrastructural-Changes in Salivary- Glands of Tsetse, Glossina-Morsitans-Morsitans, Infected with Virus and Rickettsia-Like Organisms", Journal of invertebrate pathology, vol. 56, no. 3, pp. 337-346. 5. Burand, J.P. The Sexually Transmitted Insect Virus, Hz-2V. 2009. Virologica Sinica. 24:428- 435. 6. Rallis, C.P. and J.P. Burand. 2002. Pathology and ultrastructure of the insect virus, Hz-2V, infecting agonadal female corn earworms, Helicoverpa zea . J. Invertebr. Pathol. 81:33- 44. 7. Burand, J.P., C. P. Rallis and W. Tan. 2004. Horizontal transmission of Hz-2V by virus infected Helicoverpa zea moths. J. Invertebr. Pathol. 85:128-131. 8. Abd-Alla, A.M.M., J.M. Vlak, M. Bergoin, J.E. Maruniak, A.G. Parker, J.P. Burand, J.A. Jehle and D.G. Boucias. 2009. Hytrosaviridae : a proposal for classification and nomenclature of a new insect virus family. Archives of Virology. 154:909-918. 9. Abd-Alla, A. M. M., F. Cousserans, A. G. Parker, J. A. Jehle, N. J. Parker, J. M. Vlak, A. S. Robinson, and M. Bergoin. 2008. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. Journal of Virology 82:4595-611. 10. Wang, Y., J. P. Burand and J. A. Jehle. 2007. Nudivirus genomics: diversity and classification. Virologica Sinica 22:128-136. 11. Carthew, R., Sontheimer, E., 2009. Origins and Mechanisms of miRNAs and siRNAs. Cell. 136, 642-655. 12. Bartel, D., 2009. MicroRNAs: Target Recognition and Regulatory Functions. Cell. 136, 215- 233. 13. Berezikov, E., 2011. Evolution of microRNA diversity and regulation in animals. Nature reviews Genetics. 12, 846-860. 14. Ketting, R., 2011. The many faces of RNAi. Developmental Cell. 20, 148-161. Page 138

15. Terenius, O., Papanicolaou, A., Garbutt, J.S., Eleftherianos, I., Huvenne, H., et al., 2011. RNA interference in Lepidoptera: An overview of successful and unsuccessful studies and implications for experimental design RID A-1976-2008. J. Insect Physiol. 57, 231-245. 16. Belles, X., 2010. Beyond Drosophila : RNAi In Vivo and Functional Genomics in Insects. Annu. Rev. Entomol. 55, 111-128. 17. Price, D.R.G., Gatehouse, J.A., 2008. RNAi-mediated crop protection against insects. Trends Biotechnol. 26, 393-400. 18. Whyard, S., Singh, A., Wong, S., 2009. Ingested double-stranded RNAs can act as species- specific insecticides. Insect Biochem. Mol. Biol. 39, 824-832. 19. Baum, J., Bogaert, T., Clinton, W., Heck, G., Feldmann, P., 2007. Control of coleopteran insect pests through RNA interference. Nat. Biotechnol. 25, 1322-1326. 20. Hunter, W., Ellis J., vanEngelsdorp, D., Hayes, J., Westervelt. D., Glick, E., Williams, M., Sela, I., Maori, E., Pettis, J., Cox-Foster, D., Paldi, N., 2010. Large-scale field application of RNAi technology reducing Israeli Acute Paralysis Virus disease in honey bees ( Apis mellifera , Hymenoptera: Apidae). PLoS Pathog. 6(12), e1001160 21. Abd-Alla, A.M.M., Parker, A., Bergoin, M., 2011. Tsetse salivary gland hypertrophy virus: Hope or hindrance for Tsetse control?. PLoS Neglected Tropical Diseases. 5, e1220.

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Structure, protein composition, morphogenesis and cytopathology of Glossina pallidipes salivary gland hypertrophy virus to develop rationally-designed immuno-intervention strategies to prevent virus transmission in tsetse fly colonies

Henry M. Kariithi 1, 2 , Jan W.M. van Lent 1, Adly M. M. Abd-Alla 2, Monique M. van Oers 1 and Just M. Vlak 1

1Laboratory of Virology, Wageningen University, Wageningen, The Netherlands; 2 Insect Pest Control Laboratory, International Atomic Energy Agency, Vienna, Austria

The repertoire of proteins comprising intact virions of Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) is likely to have significant consequences for virus morphology and morphogenesis. In this study, cryo-tomography of GpSGHV from hypertrophied salivary glands (SGs) of Glossina pallidipes revealed that the virion envelope and nucleocapsid components are separated by a jacket of proteinaceous material. We concluded that this material represents the GpSGHV tegument. The entire lengths of mature, extracellular virions consisted of well-organized spiral-like ridges containing regular proteinaceous substructures. Further, while nucleocapsids were restricted to the nuclei, enveloped virions were essentially located in the cell cytoplasm. This result, coupled to observation of nucleocapsids passing out of the nucleus via the nucleopore, suggests that GpSGHV nucleocapsids are assembled in the nucleus and egress to the cytoplasm of infected SG cell where virion envelopment is orchestrated. It seems that the entire envelopment occurs in the cytoplasm of infected cells and that the virions are released into the salivary gland by cell rupture or disintegration.

Since the GpSGHV virions were highly unstable in solution, we have amended the original purification procedure (Kariithi et al., 2012) and arrived at a highly purified preparation of intact virions as evidence by electron microscopy. These virions were fractionated into nucleocapsids and envelopes by two rounds of 1% Nonidet P-40-treatment, followed by 12% SDS-PAGE separation and subsequent protein identification by tandem mass spectrometry (LC-MS/MS) or defined antibodies. Fifty-four distinct, virally-encoded proteins were reliably identified by ≥ 2 peptides per protein, of which at least one was unique. We assigned 10 of these proteins to the envelope, 15 to the nucleocapsid and 29 to the tegument. In contrast to what has been found previously (Kariithi et al. , 2010) the product of orf10 , a 130 kDa protein is now found in the tegument and not in the envelope. As expected the peroral infectivity factors (PIFs) were exclusively found in the membrane fraction of the virion. Analyses of posttranslational modifications revealed that that several of the virion proteins are serine-/threonine-phosphorylated and that the proteins encoded by ORFs 38, 39, 40, 62 and 97 are O-glycosylated. Page 140

Additionally, 56 virion-associated (cellular) proteins were identified, of which the presence of α-tubulin, γ-actin and ubiquitin was verified by a proteinase K protection assay and immunoblotting. Bioinformatics analyses of the identified virion-associated GpSGHV protein sequences revealed motifs with functional and structural implications on the virus morphogenesis. Among these is the involvement of the endoplasmic reticulum and Golgi in the morphogenesis of the virus, which is compatible with the cytoplasmic assembly of GpSGHV.

Our findings provide new insights into GpSGHV morphology and morphogenesis, and point us towards areas of the virus pathogenesis in our future studies. Furthermore, the proteomic analysis may call for re-evaluation of the immuno-intervention strategy to control GpSGHV infection in tsetse fly rearing facilities for the sterile insect technique by using the newly identified envelope proteins as a basis for antibody development and GpSGHV intervention studies (Kariithi et al., 2010).

The above research were the result of a previous effort in determining the structure and sequence of the GpSGHV genome (Abd-Alla et al., 2008) and the comparison with the related hytrosavirus, Musca domestica SGHV (MdSGHV) (Garcia-Maruniak et al., 2009). This not only allowed the identification and gene assignment of virion proteins on the viral genome, but also the establishment of a new virus family Hytrosaviridae (Abd-Alla et al, 2009), which is now accepted by the Inernational Committee on Taxonomy of Viruses.

To successfully execute virus intervention studies it was necessary to study the genetic diversity of GpSGHV in African tsetse flies. From these studies it was concluded that the genetic variation was minimal (less than 1%). This implies that the virus intervention studies using antibodies are in principle generally applicable in mass rearings of tsetse.

The general conclusion is that immune intervention to block transmission of GpSGHV is generally applicable, but requires fine-tuning and testing beyond the laboratory scale.

References

Abd-Alla, A., F. Cousserans, A. Parker, J.A. Jehle, N. Parker, J.M. Vlak, A. Robinson and M. Bergoin. 2008. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. Journal of Virology 82: 4595-4611

Garcia-Maruniak, A., A.M.M. Abd-Alla, T.Z. Salem, A.G. Parker, V-U. Lietze, M.M. van Oers, J.E. Maruniak, W. Kim, J.P. Burand, F. Cousserans, A.S. Robinson, J.M. Vlak, M. Bergoin and D.G. Boucias. 2009. Two viruses that cause salivary gland hypertrophy in Glossina pallidipes Page 141

and Musca domestica are closely related and form a distinct phylogenetic clade. Journal of General Virology 90: 334 - 346.

Abd-Alla, A M.M., J.M. Vlak, M. Bergoin, J. Maruniak, A. Parker, J.P. Burand, J.A. Jehle and D.G. Boucias. 2009. Hytrosaviridae : a proposal for classification and nomenclature of a new insect virus family. Archives of Virology 154: 909-918.

Henry M. Kariithi, H.M., , I.A. Ince, S. Boeren, J. Vervoort, M. Bergoin, M.M. van Oers, A.M.M. Abd-Alla and J.M. Vlak. 2010. Proteomic analysis of Glossina pallidipes Salivary Gland Hypertrophy Virus virions for immune intervention in tsetse fly colonies. Journal of General Virology 91: 2985-2993.

Kariithi, H.M., I.A. Ince, S.A. Boeren, A.M.M. Abd-Alla, Andrew G. Parker, J.M. Vlak, S. Aksoy, J.M. Vlak and M.M. van Oers. 2011. The salivary secretome of the tseste fly Glossina pallipides (Diptera: Glossinidae) infected by salivary gland hypertrophy virus. PLoS Neglected Tropical Diseases 5(11): e1371.

Kariithi, H.M., M. Ahmadi, A.G. Parker, A.S. Robinson, G. Franz, V. Ros, I. Haq, A.M. Elashry, J.M. Vlak, M. Bergoin, J.B. Vreijsen and A.M.M. Abd-Alla. 2012. Prevalence and genetic variation of salivary gland hypertrophy virus in the wild populations of the tsetse fly Glossina pallipides (Diptera: Glossinidae) from southern and eastern Africa. Journal of Invertebrate Pathology, to be revised.

Abd-Alla, A.M.M., M. Bergoin, A.G. Parker, N.K. Maniania, J.M. Vlak, K. Bourtzis, D.G. Boucias and S. Aksoy. 2012. Improving SIT for tsetse flies through research on their symbionts and pathogens. Journal of Invertebrate Pathology, to be revised.

Kariithi, H.M., I.A. Ince, M.M. van Oers, A.M.M. Abd-Alla and J.M. Vlak. 2012. Proteomic footprints of a member of Glossinavirus (Hytrosaviridae ): An expeditious approach to virus control strategies in tsetse factories. Journal of Invertebrate Pathology, to be revised. Page 142

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FINAL RESEARCH CO-ORDINATION MEETING

JOINT FAO/IAEA DIVISION OF NUCLEAR TECHNIQUES IN FOOD AND AGRICULTURE

“IMPROVING SIT FOR TSETSE FLIES THROUGH RESEARCH ON THEIR SYMBIONTS AND PATHOGENS”

VIENNA, AUSTRIA

26-30 MARCH, 2012

“Tsetse-Wolbachia Symbiosis: from fundamental research to novel symbiont- based pest and disease control strategies”

Vangelis Doudoumis 1, George Tsiamis 1, Florence Wamwiri 2,6 , Corey Brelsfoard 2,3 , Uzma Alam 2, Emre Aksoy 2, Stelios Dalaperas 1, Alexander P. Egyir-Yawson 4, Imna Malele 5, Johnson Ouma 6, Peter Takac 7, Adly Abd-Alla 8, Serap Aksoy 2 and Kostas Bourtzis 1,9,*

1 Department of Environmental and Natural Resources Management, University of Western Greece, 2 Seferi St, 30100 Agrinio, Greece 2 Yale University School of Public Health, 60 College St., 811 LEPH, New Haven, CT 06520 USA 3 Current address: Department of Entomology, University of Kentucky, S-225 Ag. Science Center North, Lexington, KY 40546 USA 4 Vector Genetics Laboratory, Department of Animal Science, Biotechnology & Nuclear Agriculture Research Institute, Ghana Atomic Energy Commission, P.O. Box LG 80, Legon, Accra, Ghana 5 Tsetse & Trypanosomiasis Research Institute (TTRI), Majani Mapana, Off Korogwe Road, P. O. Box 1026 Tanga, Tanzania 6 Trypanosomiasis Research Centre, Kenya Agricultural Research Institute, P.O. Box 362, Kikuyu 00902 Kenya

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7 Institute of Zoology, Department of Entomology, Slovak Academy of Science, Dubravska cesta 9, 845 06 Bratislava, Slovakia 8 Insect Pest Control Laboratory, Joint FAO/IAEA Division of Nuclear Techniques in Food and Agriculture, Vienna, Austria 9 Biomedical Sciences Research Center Al. Fleming, 16672 Vari, Greece

* Author for correspondence: Kostas Bourtzis; e-mail: [email protected] Tel: +30-26410-74114, Fax: +30-26410-74171

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Abstract

Wolbachia is a widespread and highly diverse group of bacterial symbionts infecting arthropod and filarial nematode species. These alphaproteobacteria are frequently found as intracellular and maternally inherited symbionts in insect species, whose reproductive properties they manipulate by inducing cytoplasmic incompatibility (CI), parthenogenesis, male-killing and feminization, thus spreading efficiently and rapidly into host populations. Due to their unique biology, Wolbachia symbiotic associations are currently being considered as promising tools for the control of agricultural pests and medical disease vectors, including Glossina sp., which transmit African trypanosomes, the causative agents of the sleeping sickness in humans and of nagana in animals. In the present study, we report on recent studies on the detection and characterization of Wolbachia infections in Glossina species, the horizontal transfer of Wolbachia genes to tsetse chromosomes and also on how Wolbachia symbiosis could be harnessed for the development of novel symbiont-based pest and disease control strategies.

Keywords

Glossina , Wolbachia, insect symbiosis, Sodalis, Wigglesworthia , paratransgenesis, Sterile Insect Technique, Incompatible Insect Technique, Trypanosoma , African trypanosomosis, sleeping sickness, nagana, salivary gland hypertrophy virus

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Introduction

Wolbachia is an obligatory intracellular and maternally transmitted endosymbiont which has been associated with the induction of a variety of reproductive abnormalities such as cytoplasmic incompatibility (CI), male-killing, feminization, and parthenogenesis. Wolbachia symbiosis and Wolbachia -induced reproductive alterations have been reported in numerous arthropods hosts, they are, however, most common in insects (Hilgenboecker et al., 2008; Saridaki and Bourtzis, 2010; Werren et al., 2008).

Tsetse flies (Glossina spp.) are the sole vectors of Trypanosoma spp., the causative agents of sleeping sickness in humans (human African trypanosomosis, HAT) and of nagana (African animal trypanosomosis, AAT) in livestock (Leak, 1998; Welburn et al., 2001). Although epidemics have significantly declined during the last years, still tens of millions of people in Africa continue to be at risk of contracting sleeping sickness (Aksoy, 2011). There are several accepted environment-friendly methods to control the insect vector, including the sterile insect technique (SIT) (Vreysen et al., 2000).

Recently, there is an increasing interest to develop and use Wolbachia -based environment-friendly strategies for the control of insect pests and disease vectors, including tsetse flies and trypanosome transmission (Alam et al. 2011; Apostolaki et al., 2011; Bourtzis, 2008; Bourtzis and Robinson, 2006; Brelsfoard and Dobson, 2009; Brelsfoard and Dobson, 2011; Xi et al., 2005; Zabalou et al., 2004, 2009).

The main goals of our research activities in the frame of this CRP were: (a) the detection and characterization of Wolbachia infections in both laboratory and natural populations of tsetse flies and (b) investigate the possible correlation between Wolbachia and salivary gland hypertrophy virus infections. In this report, we also discuss how Wolbachia symbiosis could be harnessed for the control of tsetse flies and trypanosomosis.

Detection of Wolbachia in laboratory and natural populations of tsetse flies Page 147

We applied a specific 16S rRNA PCR assay (Doudoumis et al., 2012) to detect the presence of Wolbachia in a total of 5339 specimens of eleven different Glossina species (G. m. morsitans, G. m. centralis, G. m. submorsitans, G. austeni, G. brevipalpis, G. pallidipes, G. p. palpalis, G. p. gambiensis, G. fuscipes fuscipes, G. tachinoides and G. medicorum ). These specimens consisted of eight laboratory stocks (Table 1) and 13 natural populations originating from 13 African countries (Table 2). The different laboratory stocks were (FAO/IAEA-Seibersdorf, Yale University-EPH, SAS-Bratislava, KARI-TRC, CIRDES, Antwerp, Pangani/Tanga Tanzania, UGA/IAEA). While the natural populations were from (Tanzania, South Africa, Zambia, Zimbabwe, Kenya, Senegal, Guinea, Ethiopia, Uganda, Democratic Republic of Congo, Burkina Faso, Ghana and Mali).

Characterization of Wolbachia strains from tsetse flies

The characterization of tsetse Wolbachia strains was based on Multi Locus Typing System (MLST) and the wsp gene (outer surface protein). We selected a total of 23 Wolbachia -infected Glossina specimens from different areas and species to be genotyped by amplifying the five MLST housekeeping genes ( gatB, coxA, hcpA, ftsZ and fbpA ), as well as the wsp gene (Baldo et al. 2006).

Specifically, we characterized Wolbachia infections of eight different Glossina species from five laboratory lines and 18 natural populations originating from nine African countries. So far, 12 allelic profiles or Sequence Types (ST) were detected in tsetse flies Wolbachia strains; all of them were new STs, based on the available data in the Wolbachia MLST database (Table 3).

Moreover, the same 23 samples were genotyped using the wsp (Wolbachia surface protein) gene, for which 15 alleles were found. We also identified their WSP HVR profile (a combination of the four HVR amino acid haplotypes) as described in Baldo et al. 2006. To date, a total of fourteen WSP HVR profiles were found, of which twelve were new entries to the Wolbachia WSP database (Table 4).

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Phylogenetic analysis

All sequences generated in the frame of this study were manually edited with SeqManII (DNAStar), adjusted by eye, aligned using MUSCLE (Egdar, 2004) and used for the construction of Neighbour–Joining (NJ) phylogenetic trees, according to the method of Jukes and Cantor as genetic distance model, with 1000 replicates, as implemented in Geneius v5.3.4 [http://www.geneious.com/ ]. Four phylogenetic trees are presented in this report based on: a) the concatenated data set of the MLST Wolbachia genes, b) the Wolbachia wsp gene, c) the Wolbachia 16S rRNA gene and d) the mitochondrial cytochrome oxidase subunit I of the Glossina species studied for the presence of Wolbachia (Figures 1-4 respectively).

Horizontal transfer of Wolbachia genes to tsetse chromosomes

During the 16S rRNA screening and the MLST analysis, evidence for horizontal transfer of the 16S rRNA, gatB, coxA, hcpA, ftsZ, fbpA and wsp genes to the host chromosomes was observed in specimens of three Glossina species ( G. morsitans morsitans, G. pallidipes and G. austeni ). Cloning and sequencing analysis showed that the horizontally transferred Wolbachia genes were pseudogenized through the accumulation of deletion? events. It is interesting to note that different pseudogenization events were detected for most of the genes (Figures 5-10).

Wolbachia genomics – Wolbachia ankyrins

Since the sequencing of the G. m. morsitans genome was in progress (IGGI: International Glossina Genome Initiative), we tried to “fish out” Wolbachia gene sequences. A single tetracycline-treated female fly was used for the genome project which means that it was free of any cytoplasmic Wolbachia. So, any Wolbachia sequences present in the tsetse genomic data should stem from chromosomally inserted Wolbachia . Sanger, 454 and Solexa technologies were used for the genome project. Tools that were used to “fish out” Wolbachia-specific sequences included: (a) Page 149

MIRA and (b) AMOS. As a reference sequence, we initially used the available Wolbachia complete genomes from the strains wRi, wMel, wPip and wBm. The genome of the cytoplasmic wGmm strain, sequenced by Prof. Serap Aksoy’s lab by 454 technology, was also included. In total. 0.3% of the tse-tse whole genome sequence data analyzed were Wolbachia sequences. The assembly resulted in 314 contigs with a total number of 675,602 bp. There were only 10 (>15,000 /10) large size contigs, while the small sized ones numbered 304. Number of CDSsFs: >400. Preliminary sequence and FISH analysis showed that at least two HGT events may have occurred, which should be separated before attempting annotation. The separation was initially based on a “sequence signature” approach to pool “strain specific sequence data” using ClaMS and later on a SNP-based approach using AMOS. The later approach was successful resulting in the isolation of the two insertion events, their assembly and annotation.

The annotation was done with Glimmer, Blast and Artemis. The three tsetse Wolbachia types (the cytplasmic and the two insertions) were compared to wMel and wRi, two other A-supergroup CI-inducing Wolbachia strains. The wGmm strain seems to be more closely related to the Wolbachia strain wMel than wRi. The genome size of the tsetse Wolbachia chromosomal insertions is reduced and has fewer CDSs, shorter genes, an increased number of pseudogenes for the chromosomal insertions, as expected. The GC% content is the same etc. The majority of the chromosomal CDSs are pseudogenes, they contain a lot of SNPs, as well as several deletions or insertions, which render these CDSs inactive. Several characteristic features of the Wolbachia genome are retained, such as the presence of ankyrin genes and [prophages, ISs etc].

It has been shown that ankyrins mediate protein-protein interactions and act as transcriptional and developmental regulators as well as inhibitors and toxins (Bork 1993, Michaely & Bennett 1992, Sedgick & Smerdon 1999, Mosavi et al. 2004). Ankyrins are widespread among eukaryotes while they seem to be more restricted in prokaryotes. However, a few bacteria contain a significant number of ankyrin genes including the CI-inducing Wolbachia strains of wMel, wRi and wPip (Seshadri et al. 2003; Wu et al. 2004; Ogata et al. 2005, 2006; Klasson et al. 2008, 2009). Ankyrins were also detected in both the cytoplasmic wGmm strain and the chromosomal

Page 150 insertions; the later have essentially been pseudogenized and are probably not expressed.

Wolbachia and salivary gland hypertrophy virus

The specimens were also screened for the presence of the tsetse salivary gland hypertrophy virus (Gp -SGHV). This screen suggests that there is a negative correlation between the presence of Wolbachia and SGHV (Table 5, Figures 11-12).

Wolbachia -induced cytoplasmic incompatibility - Wolbachia -based control strategies

Wolbachia -induced CI has been proposed as a potential mechanism for the control of agricultural pests and disease vectors (Beard et al., 1998; Beard et al., 1993a, b; Bourtzis, 2008; Bourtzis and O’Neill, 1998; Dobson, 2003; Sinkins and Godfray, 2004). Our study shows that Wolbachia is present in both laboratory and natural populations of tsetse flies, while previous studies have found that the symbiont is mainly present in the reproductive tissues of tsetse (Aksoy, 2000; Cheng et al., 2000; Doudoumis et al., 2012; O’Neill et al., 1993), Alam and colleagues (2011) recently reported that the presence of Wolbachia is indeed responsible for the induction of strong CI in G. m. morsitans rendering any similar experiments, like the transfer of tsetse Wolbachia strains in Drosophila species, unnecessary (Alam et al., 2011).

Expression of strong CI in Wolbachia -infected G. m. morsitans opens new routes for population control of the insect vector and of trypanosomosis. This can be done with three potential approaches: (a) to use Wolbachia -induced CI as a population suppression mechanism in a way analogous to SIT (Apostolaki et al., 2011; Laven, 1967; Zabalou et al., 2009; Zabalou et al., 2004); (b) to use Wolbachia -induced CI as a spreading/replacement mechanism for desired phenotypes. Infected females have a reproductive advantage over uninfected females, since they can mate with both uninfected and infected males, thus their genotype invades populations in nature (Bourtzis, 2008; Bourtzis and Robinson, 2006; Bourtzis and O’Neill, 1998; Brelsfoard and Dobson, 2009; Dobson et al., 2002; Rasgon, 2007, 2008; Sinkins and Gould, Page 151

2006) and (c) Wolbachia -induced CI can also be used to drive trypanosome resistant paratransgenic tsetse into natural populations to replace their parasite-susceptible counterparts (Alam et al., 2011). Based on the fact that all there tsetse symbionts are maternally transmitted to the progeny, resistance genes expressed by Sodalis can be propagated by Wolbachia induced CI , if complete transmission of Sodalis and Wolbachia can be ensured (Aksoy et al., 2008; Aksoy and Weiss, 2007; Rio et al., 2004).

References

Aksoy, S., 2000. Tsetse--A haven for microorganisms. Parasitol Today. 16 , 114-8. Aksoy, S., 2011. Sleeping sickness elimination in sight: time to celebrate and reflect, but not relax. PLoS Negl Trop Dis. 5 , e1008. Aksoy, S., Weiss, B. L., Symbiosis-Based Technological Advances to Improve Tsetse Glossina spp . SIT Application. In: M. J. B. Vreysen, et al., Eds.), Area-Wide Control of Insect Pests. Springer, Dordrecht, The Netherlands, 2007, pp. 137- 148. Aksoy, S., Weiss, B., Attardo, G., 2008. Paratransgenesis applied for control of tsetse transmitted sleeping sickness. Adv Exp Med Biol. 627 , 35-48. Alam, U., Medlock, J., Brelsfoard, C., Pais, R., Lohs, C., Balmand, S., Carnogursky, J., Heddi, A., Takac, P., Galvani, A., Aksoy, S., Wolbachia symbiont infections induce strong cytoplasmic incompatibility in the tsetse fly Glossina morsitans . PLoS Pathogens, 2011 Dec;7(12):e1002415. Epub 2011 Dec 8. Apostolaki, Α., Livadaras, I., Saridaki, A., Chrysargyris, A., Savakis, C., Bourtzis, K., 2011. Transinfection of the olive fruit fly Bactrocera oleae with Wolbachia : towards a symbiont-based population control strategy. Journal of Applied Entomology 135 , 546-553. Baldo L, Hotopp JCD, Jolley KA, et al. (2006) Multilocus sequence typing system for the endosymbiont Wolbachia pipientis . Appl. Environ. Microbiol. 72 :7098- 7110. Beard, C. B., O'Neill, S. L., Mason, P., Mandelco, L., Woese, C. R., Tesh, R. B., Richards, F. F., Aksoy, S., 1993a. Genetic transformation and phylogeny of bacterial symbionts from tsetse. Insect Mol Biol. 1, 123-31.

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Table 1 : Wolbachia prevalence in laboratory colonies and field populations of 11 Glossina species (Numbers in parentheses indicate the number of Wolbachia –infected adult flies / total number of flies tested)

Glossina species Field % Lab % Total % G. m. centralis Not analyzed (3/3) 100.0% (3/3) 100.0% G. austeni (208/235) 88.5% Not analyzed (208/235) 88.5% G. m. morsitans (521/604) 86.3% (109/109) 100.0% (630/713) 88.4% G. medicorum (20/94) 21.3% Not analyzed (20/94) 21.3% G. brevipalpis (3/52) 5.8% (14/39) 35.9% (17/91) 18.7% G. p. palpalis (32/299) 10.7% (0/36) 0.0% (32/335) 9.6% G. tachinoides (37/433) 8.5% (0/7) 0.0% (37/440) 8.4% G. m. submorsitans (6/142) 4.2% Not analyzed (6/142) 4.2% G. p. gambiensis (28/1194) 2.3% (0/89) 0.0% (28/1283) 2.2% G. pallidipes (29/1672) 1.7% (8/242) 3.3% (37/1914) 1.9% G. f. fuscipes (0/53) 0.0% (0/36) 0.0% (0/89) 0.0%

TOTAL (884/4778) 18.5% (134/561) 23.9% (1018/5339) 19.1%

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Table 2 : Wolbachia prevalence in eight different laboratory lines and in natural populations originating from 13 African countries (Numbers in parentheses indicate the number of Wolbachia –infected adult flies / total number of flies tested)

Glossina species Country (Area, Collection Date) Prevalence Tanzania (Jozani, 1997) (22/42) 52.4% Tanzania (Zanzibar, 1995) (75/78) 96.2% G. austeni South Africa (Zululand, 1999) (79/83) 95.2% Kenya (Shimba Hills, 2010) (30/30) 100.0% Coastal Tanzania (Muhoro, Rufiji Tanzania) (2/2) 100.0% Seibersdorf lab-colony (1995) (14/34) 41.2% Lab (Coastal Tanzania-Pangani, Tanga Tanzania) (0/5) 0.0% G. brevipalpis South Africa (Zululand, 1995) (1/50) 2.0% Coastal Tanzania (Muhoro, Rufiji Tanzania) (2/2) 100.0% Seibersdorf lab-colony (1995) (0/36) 0.0% G. f. fuscipes Uganda (Buvuma island, 1994) (0/53) 0.0% G. medicorum Burkina Faso (Comoe, 2008) (20/94) 21.3% G. m. centralis Yale lab-colony (2008) (3/3) 100.0% Burkina Faso (Nazinga, 2009) (0/3) 0.0% G. m. submorsitans Burkina Faso (Comoe Folonzo, 2007) (2/30) 6.7% Burkina Faso (Comoe, 2008) (4/109) 3.7% Seibersdorf lab-colony (1995) (0/36) 0.0% G. p. palpalis Democratic Republic of Congo (Zaire, 1995) (0/48) 0.0% Ghana (32/251) 12.7% Seibersdorf lab-colony (1995) (0/7) 0.0% Burkina Faso (Nazinga, 2009) (0/15) 0.0% Burkina Faso (Comoe Folonzo, 2007) (5/112) 4.5% G. tachinoides Burkina Faso (Comoe, 2008) (0/72) 0.0% Ghana (Pong Tamale, Walewale, 2008) ( 5/46) 10.9% Ghana (Walewale, 2008) (27/149) 18.1% Ghana (Fumbissi, 2008) (0/39) 0.0% Coastal Tanzania (Utete, Rufiji Tanzania) (2/3) 66.7% G. m. morsitans Zambia (MFWE, Eastern Zambia, 2007) (122/122) 100.0%

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KARI-TRC lab-colony (2008) (89/89) 100.0% Tanzania (Ruma, 2005) (100/100) 100.0% Zimbabwe (Gokwe, 2006) (7/74) 9.5% Zimbabwe (Kemukura, 2006) (26/26) 100.0% Zimbabwe (M.Chiuy, 1994) (33/36) 91.7% Zimbabwe (Makuti, 2006) (95/99) 96.0% Zimbabwe (Mukond, 1994) (35/36) 97.2% Zimbabwe (Mushumb, 2006) (3/8) 37.5% Zimbabwe (Rukomeshi, 2006) (98/100) 98.0% Yale lab-colony (2008) (5/5) 100.0% Antwerp lab-colony (2010) (10/10) 100.0% Bratislava lab-colony (2010) (5/5) 100.0% Zambia (MFWE, Eastern Zambia, 2007) (5/203) 2.5% KARI-TRC lab-colony (2008) (3/99) 3.0% Kenya (Mewa, Katotoi and Meru national park, 2007) (0/470) 0.0% Ethiopia (Arba Minch, 2007) (2/454) 0.4% Seibersdorf lab-colony (2008) (0/138) 0.0% Tanzania (Ruma, 2005) (3/83) 3.6% Tanzania (Mlembuli and Tunguli, 2009) (0/94) 0.0% Zimbabwe (Mushumb, 2006) (0/50) 0.0% G. pallidipes Zimbabwe (Gokwe, 2006) (0/150) 0.0% Zimbabwe (Rukomeshi, 2006) (5/59) 8.5% Zimbabwe (Makuti, 2006) (4/96) 4.2% Lab (Mainland Uganda, UGA /IAEA) (5/5) 100.0% Mainland Tanzania (Death Valley Tanzania,Naitolia (4/6) 66.7% Arusha) Coastal Tanzania (Muhoro, Rufiji Tanzania) (3/4) 75.0% Coastal Tanzania (Muyuyu, Rufiji Tanzania) (3/3) 100.0% CIRDES lab-colony (1995) (0/32) 0.0% CIRDES lab-colony (2005) (0/57) 0.0% Senegal (Diacksao Peul and Pout, 2009) (1/188) 0.5% G. p. gambiensis Guinea (Kansaba, Mini Pontda, Kindoya and Ghada (0/180) 0.0% Oundou, 2009) Guinea (Alahine, 2009) (0/29) 0.0% Page 159

Guinea (Boureya Kolonko, 2009) (0/36) 0.0% Guinea (Fefe, 2009) (0/29) 0.0% Guinea (Kansaba, 2009) (0/19) 0.0% Guinea (Kindoya, 2009) (1/12) 8.3% Guinea (Lemonako, 2009) (0/30) 0.0% Guinea (Togoue, 2009) (0/32) 0.0% Guinea (Conakry, 2010) (5/138) 3.6% Burkina Faso (Comoe, 2008) (0/12) 0.0% Burkina Faso (Comoe Folonzo, 2007) (1/53) 1.9% Burkina Faso (Kenedougou, 2007) (1/37) 2.7% Burkina Faso (Houet Bama, 2007) (1/69) 1.4% Guinea (Fefe, Togoue, Alahine, Boureya Kolonko, (5/94) 5.3% 2009-2010) Guinea ( Boureya Kolonko, Kansaba, Kindoya, Ghada (3/94) 3.2% Oundou, 2009-2010) Mali (Fijira, 2009) (0/14) 0.0% Senegal (Diaka Madia, 2009) (0/42) 0.0% Senegal (Tambacounda, 2008) (3/38) 7.9% Senegal (Simenti, 2008) (6/33) 18.2% Senegal (Kédougou, 2008) (1/15) 6.7%

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Table 3: MLST allele profiles of the Wolbachia strains from 23 Glossina populations.

Country Wolbachia MLST Code Species (Area, Collection Date) ST gatB coxA hcpA ftsZ fbpA GmcY G. m. centralis Yale lab-colony (2008) 226 141 127 23 114 15 12.3A G. m. morsitans Zambia (MFWE, Eastern Zambia, 2007) 226 141 127 23 114 15 32.3D G. m. morsitans Zimbabwe (Makuti, 2006) 226 141 127 23 114 15 30.9D G. m. morsitans Zimbabwe (Rukomeshi, 2006) 227 141 127 23 115 15 34.7G G. m. morsitans Tanzania (Ruma, 2005) N/C 141 128 23 114 N/A GmmY G. m. morsitans Yale lab-colony (2008) 228 8 127 23 113 15 24.4A G. m. morsitans KARI-TRC lab-colony (2008) 229 142 128 23 113 15 24.1A G. pallidipes KARI-TRC lab colony (2008) New New 127 New 113 15 10.10E G. pallidipes Zambia (MFWE, Eastern Zambia, 2007) N/C New 128 New New N/A 32.11G G. pallidipes Zimbabwe (Makuti, 2006) N/C 141 128 23 New N/A 30.10G G. pallidipes Zimbabwe (Rukomeshi, 2006) N/C 141 128 23 114 N/A 34.4A G. pallidipes Tanzania (Ruma, 2005) N/C 141 128 23 New N/A 15.5B G. pallidipes Ethiopia (Arba Minch, 2007) 232 144 47 149 116 202 05.2B G. austeni South Africa (Zululand, 1999) 231 128 109 127 98 20 013.11B G. austeni Tanzania (Zanzibar, 1995) 231 128 109 127 98 20 015.10B G. austeni Tanzania (Jozani, 1997, F) 231 128 109 127 98 20 015.1B G. austeni Tanzania (Zanzibar, 1995) 231 128 109 127 98 20 GauK G. austeni Kenya (Shimba Hills, 2010) 197 128 108 127 98 20 405.11F G. p. gambiensis Guinea (Kindoya, 2009) 233 145 130 150 117 203 184.Gpp G. p. palpalis Ghana (NA, NA) New New New New New New 011.4H G. brevipalpis South Africa (Zululand, 1995) New 14 129 New 56 15 09.7G G. brevipalpis Seibersdorf lab-colony (1995) 230 143 129 23 56 15 523.4F G. medicorum Burkina Fasso (Comoe, 2008) New New New New New New

New: new allele or ST in the Wolbachia MLST database. N/C: not completed, in progress. N/A: not amplified, failure to detect amplification product with standard primers (new primers are currently under evaluation).

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Table 4: WSP HVR profiles of the Wolbachia strains infecting 23 Glossina populations.

Code Species Country (Area, Collection Date) wsp HVR1 HVR2 HVR3 HVR4 GmcY G. m. centralis Yale lab-colony (2008) 550 193 9 221 202 12.3A G. m. morsitans Zambia (MFWE, Eastern Zambia, 2007) 548 192 9 12 202 GmmY G. m. morsitans Yale lab-colony (2008) 548 192 9 12 202 30.10G G. pallidipes Zimbabwe (Rukomeshi, 2006) 548 192 9 12 202 34.7G G. m. morsitans Tanzania (Ruma, 2005) New* 192 9 15 202 34.4A G. pallidipes Tanzania (Ruma, 2005) New* 192 9 15 202 32.3D G. m. morsitans Zimbabwe (Makuti, 2006) 356 142 9 12 9 30.9D G. m. morsitans Zimbabwe (Rukomeshi, 2006) 356 142 9 12 9 32.11G G. pallidipes Zimbabwe (Makuti, 2006) New New 9 12 9 24.1A G. pallidipes KARI-TRC lab colony (2008) New New 9 12 9 24.4A G. m. morsitans KARI-TRC lab-colony (2008) 549 142 9 223 9 15.5B G. pallidipes Ethiopia (Arba Minch, 2007) 552 195 224 224 63 09.7G G. brevipalpis Seibersdorf lab-colony (1995) 11 9 9 12 9 011.4H G. brevipalpis South Africa (Zululand, 1995) 11 9 9 12 9 013.11B G. austeni Tanzania (Zanzibar, 1995) 551 180 40 210 18 015.10B G. austeni Tanzania (Jozani, 1997, F) 551 180 40 210 18 015.1B G. austeni Tanzania (Zanzibar, 1995) 551 180 40 210 18 05.2B G. austeni South Africa (Zululand, 1999) 551 180 40 210 18 GauK G. austeni Kenya (Shimba Hills, 2010) 507 180 40 210 18 405.11F G. p. gambiensis Guinea (Kindoya, 2009) 553 194 223 222 220 184.Gpp G. p. palpalis Ghana (NA, NA) New New New 22 140 523.4F G. medicorum Burkina Faso (Comoe, 2008) New New New New New 10.10E G. pallidipes Zambia (MFWE, Eastern Zambia, 2007) New New 9 New 9

New: new allele in the Wolbachia MLST database. *: identical

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Table 5: Wolbachia (W) and SGHV (V) prevalence in laboratory lines and natural populations of tsetse flies (numbers in parentheses indicate the number of infected adult flies / total number of flies tested)

TYPE FIELD LAB TOTAL

W+ V+ 3.2% (142/4497) 0.6% (3/537) 2.9% (145/5034)

W+ V- 15.1% (680/4497) 19.9% (107/537) 15.6% (787/5034)

W- V+ 10.2% (459/4497) 16.9% (91/537) 10.9% (550/5034)

W- V- 71.5% (3216/4497) 62.6% (336/537) 70.6% (3552/5034)

W+ 18.3% (822/4497) 20.5% (110/537) 18.5% (932/5034)

V+ 13.4% (601/4497) 17.5% (94/537) 13.8% (695/5034)

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Figure 1 : Phylogeny based on the concatenated Wolbachia MLST data (2,079 bp).

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Figure 2 : Phylogeny based on the Wolbachia wsp gene sequence.

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Figure 3 : Phylogeny based on the Wolbachia 16S rRNA sequence (438bp).

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Figure 4 : Phylogeny based on mitochondrial COI gene sequences (~300bp).

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Figure 5 : Overview of fragments deleted from the Wolbachia 16S rRNA gene.

The 16S rRNA fragments from tsetse Wolbachia strains aligned with the corresponding regions of strain wMel using MUSCLE, as implemented in Geneius v5.3.4. Black lines represent the deletion region; the numbers show the positions before and after the deletions in respect to the wMel genome and the right-left red arrows below the number indicate the size of deletion in base pairs. Two types of deleted fragments were detected: a) the type 1 (T1) in seven G. m. morsitans samples (Gmtet, GmmY, 12.3A, 24.4A, 30.9D, 32.3D and 34.7G), in five G. pallidipes (10.10E, 24.1A, 30.10G, 32.11G, 34.4A) and one G.austeni (013.11B), b) the type 2 (T2) in two G. m. morsitans samples (24.4A, 32.3D) and one G. pallidipes (10.10E).

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Figure 6 : Overview of fragments deleted from the Wolbachia fbpA gene.

The fbpA fragments from tsetse Wolbachia strains, aligned with the corresponding regions of strain wMel using MUSCLE, as implemented in Geneius v5.3.4. Black lines represent the deletion region and the number indicates the size of the deletion in base pairs; the numbers show the positions before and after the deletions in respect to the wMel genome. Two types of deleted fragments were detected in several tsetse fly samples: a) the type 1 (T1) in seven G. m. morsitans samples (Gmtet, GmmY, 12.3A, 24.4A, 30.9D, 32.3D and 34.7G), in five G. pallidipes (10.10E, 24.1A, 30.10G, 32.11G, 34.4A) and one G.austeni (013.11B), b) the type 2 (T2) in one G. m. morsitans sample (32.3D) and one G. pallidipes (10.10E).

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Figure 7 : Overview of fragments deleted from the Wolbachia ftsZ gene.

ftsZ fragments from tsetse Wolbachia strains aligned with the corresponding regions of strain wMel using MUSCLE, as implemented in Geneius v5.3.4. Black lines represent the deletion region and the number indicates the size of the deletion in base pairs; the numbers show the positions before and after the deletions in respect to the wMel genome. Two types of deleted fragments were detected in several tsetse fly samples: a) the type 1 (T1) in threee G. m. morsitans samples (Gmtet, GmmY, 12.3A and 24.4A) and in three G. pallidipes (10.10E, 24.1A and 32.11G), b) the type 2 (T2) in one G. m. morsitans sample (GmmY).

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Figure 8 : Overview of pseudogenized fragments of the Wolbachia wsp gene.

wsp fragments from tsetse Wolbachia strains aligned with the corresponding regions of strain wMel using MUSCLE, as implemented in Geneius v5.3.4. Black lines represent the deletion region, and the number indicates the size of the deletion in base pairs; the numbers show the positions before and after the deletions in respect to the wMel genome. Four types of pseudogenized fragments were detected in several tsetse fly samples and additionally, an insertion of 4 nucleotides was found in type 4 between positions 1023370-75: a) the type 1 (T1) in one G. m. morsitans samples (Gmtet), b) the type 2 (T2) in two G. m. morsitans samples (30.9D and 32.3D), c) the type 3 (T3) in two G. m. morsitans samples (30.9D and 32.3D), d) the type 4 (T4) in one G. pallidipes (10.10E).

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Figure 9: Overview of fragments deleted from the Wolbachia coxA gene.

coxA fragments from tsetse Wolbachia strains aligned with the corresponding regions of strain wMel using MUSCLE, as implemented in Geneius v5.3.4. Black dashes represent the deletion region, and the number indicates the size of the deletion in base pairs; the numbers show the positions before and after the deletions in respect to the wMel genome. Only one type of deleted fragment was detected in a G. m. morsitans sample (30.9D).

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Figure 10 : Overview of fragments deleted from the Wolbachia gatB gene.

gatB fragments from tsetse Wolbachia strains aligned with the corresponding regions of strain wMel using MUSCLE, as implemented in Geneius v5.3.4. Black dashes represent the deletion region, and the number indicates the size of the deletion in base pairs; the numbers show the positions before and after the deletions in respect to the wMel genome. Two types of deleted fragments were detected in one G. m. morsitans sample (24.4A).

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Figure 11 : Wolbachia and SGHV patterns of infections including both laboratory and field samples.

Wolbachia / SGHV

100% 90% 80% 70% 60% W+ / V+ 50% Wolbachia W+ / V- 40% 30%

Positive 20% 10% 0%

s s i es tan teni id tans i s ens i s i lidipes no rs l i dicorum a e G. au ch ta bmo . brevipalpis gamb G. p m G p. G. . su G. G. m. mor G. m G. Species

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Figure 12 : Wolbachia and SGHV patterns of infections

W+ / V+ Wolbachia / SGHV W+ / V- 100% 80% 60%

40%

20%

Wolbachia 0% FIELD LAB FIELD LAB FIELD LAB FIELD NO LAB FIELD NO LAB FIELD NO LAB FIELD NO LAB FIELD NO LAB -20% G. m. morsitans G. brevipalpis G. pallidipes G. p. gambiensis G. austeni G. m. G. medicorum G. tachinoides

Positive Positive -40% submorsitans

-60%

-80%

-100% Species / Field-Lab

Page 175

Title: Ultrastructure of the Salivary Glands of Non-Infected and Infected Glands in Glossina pallidipes by the Salivary Glands Hypertrophy Virus (SGHV)

Laura Guerra 1* , John G. Stoffolano Jr 2, Gabriella Gambellini 1, Valentina Laghezza Masci 1, Maria Cristina Belardinelli 1 and Anna Maria Fausto 1

1Dipartimento per le Innovazioni dei sistemi Biologici, Agroalimentari e Forestali, Università della Tuscia, Largo dell’Università, 01100 Viterbo, Italy, 2Department of Plant, Soil and Insect Sciences, Division of Entomology, University of Massachusetts, Amherst, MA 01003, USA

*Corresponding author: Dipartimento per le Innovazioni dei sistemi Biologici, Agroalimentari e Forestali, Università della Tuscia, Largo dell’Università, 01100 Viterbo, Italy; e-mail: [email protected], phone: +390761357152, Fax: +390761357389

Abstract This study allowed us to determine the general organization of the salivary glands in Glossina pallidipes . Their morphology is characterized by three distinct regions: secretory, reabsorptive and proximal. The light microscope, the scanning and transmission electron microscopy analyses showed a detailed morphology and ultrastructure for each region, highlighting the changes in the composition and organization of tissues and cells of the entire gland. The salivary glands infected with the salivary gland hypertrophy (SGH) virus showed a severe hypertrophy of the glands, accompanied by a profound alteration of their morphology and ultrastructure. In addition, the SEM analysis clearly revealed a disruption of the muscular fibers surrounding the secretory region of the glands. The morphological changes and alterations in the muscular tissue, caused by viral infection, could be an important aspect of the pathology and may shed light on the mode of action of the SGH virus. These results are discussed in relation to the alterations produced in both the muscular and secretory aspects of infected glands and how their involvement might affect normal salivation. Finally, the potential effect of tsetse flies having the virus on their ability to transmit the trypanosome parasite is discussed.

Key words: salivary gland hypertrophy virus, Glossina pallidipes , muscular tissue, scanning electron microscopy, transmission electron microscopy, trypanosome

1. Introduction Tsetse flies ( Diptera: Glossinidae ) are obligate blood feeding insects that are important disease vectors given their involvement in the transmission of different pathogenic trypanosome species, causing human sleeping sickness and livestock trypanosomiasis in Africa. Trypanosomes of the Trypanosome brucei group, including the two human-pathogenic subspecies, T.b.gambiense and T.b.rhodesiense , have to go through a complex developmental cycle in the alimentary tract and in salivary glands of the tsetse fly (Van Den Abbeele et al., 1999). Thus, the salivary glands play an important role in the transmission of the parasite. Page 176

Many species of tsetse flies are infected with a virus that causes salivary gland hypertrophy. In particular, the Salivary Glands Hypertrophy Virus (SGHV) has been found and studied, not only in tsetse flies, but also in Musca domestica (Lietze et al., 2011). Recently, the genomic organization of SGHVs was determined and, on the basis of the available morphological, (patho)biological, genomic and phylogenetic data, it was proposed that these viruses are members of a new virus family, named Hytrosaviridae (Abd-Alla et al., 2009). These viruses, characterized by rod-shaped and large circular double stranded DNA genomes, replicate in the nucleus of salivary gland cells in adult flies, causing distinct tissue hypertrophy and reduction of host fertility (Lietze et al., 2011). Our study focused on Glossina pallidipes , where the first identification of virus particles, associated with SGH symptoms, dates to the 1970s (Jaenson, 1978). G. pallidipes Salivary Glands Hypertrophy Virus (GpSGHV), in addition to infecting the salivary glands, has been reported to replicate in the female milk gland, as well as in gonadal tissue, resulting in testicular degeneration and ovarian abnormalities (Sang et al., 1999, 1998). Electron microscopic observations of virus particles either in thin sections of hypertrophied salivary glands or from sucrose density gradient-purified, negatively stained preparations, showed enveloped bacilliform virions. The pathology of infected cells and effect on various organelles in the salivary glands has been documented (Jura et al, 1989; Kokwaro et al., 1991, 1990; Otieno et al. 1980). However, some aspects of this extremely interesting relationship between the virus and the host need further study, such as an investigation of the muscle cell layer or sheath covering the salivary glands and its cytopathology, which is caused by the viral infection. Therefore, the focus of this research was to increase our knowledge of the overall structure of both the non-infected and infected salivary glands and their morphological and ultrastructural differences, to highlight the viral effects and their possible involvement on the alterations of the glands physiology, and to better understand the mode of action of SGH virus. Furthermore, this study may shed light on the possible relationship between the structural and probably functional changes of the salivary glands and how this may affect the ability of tsetse flies in their role as vectors of trypanosomes.

2. Materials and Methods 2.1 Animals G. pallidipes pupae were received from a laboratory colony maintained in the Insect Pest Control Laboratory of the International Atomic Energy Agency (IAEA) in Vienna, Austria. These pupae were maintained at a temperature of 24°C and 70% RH, with a photoperiod of 12 hours, as already described in previous studies (Adly Abd-Alla et al. 2007; Feldmann, 1994; Gooding et al., 1997). The adult insects were reared by providing them with a meal of sugar and the average period of life observed was about 10 days. Specimens of both sexes of the adult insects were dissected taking samples of healthy salivary glands and hypertrophied glands, infected by the SGH virus.

2.2 Light microscopy Samples of salivary glands infected and non-infected were processed for light microscopy observations. In particular, light microscope sections were stained for general histology in 3% Page 177

toluidine blue and the observations were made using a computerized image analysis system, which includes a microscope Zeiss (Axiophot), equipped with a video color camera (Axio Cam MRC, Arese, Milano-Italy) and a software (KS 300and AxioVision).

2.3 Scanning Electron Microscopy (SEM) For the SEM investigation, samples of salivary glands infected and non-infected were fixed for one night in 2.5% glutaraldehyde in 0.1 M cacodylate 3% sucrose buffer pH 7.2. Subsequently, the material was washed in cacodylate buffer 0.1 M, for one night, and then, post fixed with 1% osmium tetroxide in cacodylate buffer 0.1 M. The specimens were dehydrated through a series of steps in acetone at progressively increasing concentrations (from 50% to 100%). After these steps, the material was dried with liquid carbon dioxide using the critical point dryer (Balzer Union CPD 020), mounted on a special sample holder and a metallic gold alloy evaporator Balzer Union MD 010. So treated, the samples were observed under a Jeol JSM 5200 microscope.

2.4 Transmission Electron Microscopy (TEM) For the TEM survey, samples of salivary glands infected and non-infected were fixed for one night, as described for the SEM analysis, in 2.5% glutaraldehyde in 0.1 M cacodylate 3% sucrose buffer pH 7.2. The material was washed in cacodylate buffer 0.1 M, for one night, and then, post fixed with 1% osmium tetroxide in cacodylate buffer 0.1 M. The specimens were dehydrated through a series of steps in acetone at progressively increasing concentrations (from 50% to 100%) and then, samples were infiltrated in Epon resin (TAAB, England). The ultra-thin (thickness of 60-80 nm) sections were collected on special screens of copper and were contrasted with uranyl acetate and lead citrate. Finally, the samples were observed using the Jeol JEM EX II, 120 kV, transmission electron microscope.

3. Results The salivary glands of G. pallidipes consist of two tubular structures and, in particular, the glands appear as thin and transparent tubes that extend into the abdomen on either side of the gut. In the normal and healthy salivary glands, each paired gland shows three distinct regions: secretory, reabsorptive, and proximal. This last portion continues in a duct joining a common salivary duct (Fig.1A). The external morphology of healthy salivary glands was investigated by light microscope and the SEM analysis. The secretory region extends from the abdomen into the thorax and is characterized by a thick muscle tissue with evident longitudinal muscle fibers, running throughout the distal portion of the gland (Figs. 1D,E), whereas the outer surface of the reabsorptive and proximal regions consists of a thin tube devoid of a muscular coat (Figs. 1A,B,C). The infection with SGH virus greatly enlarges the salivary glands and this hypertrophy is uniform over the entire length of the gland. In fact, each of the salivary gland pairs are equally affected, increase in diameter and appear whitish and pale and are therefore easily Page 178

distinguishable from non-infected and transparent glands (Fig. 2A). However, hypertrophy primarily affects the secretory region of the glands, which fill most of the abdominal cavity, where they become greatly entangled with the fat bodies and the tracheae (Fig. 2A). The observations of the SEM show the evident alteration of the muscular tissue in the hypertrophied salivary glands. In fact, SEM micrographs revealed that the muscle fibers lose their organization and arrangement in longitudinal fibers and, also, they are often broken with the formation of some cavities along their length (Figs. 2D,E). Figures 3 and 4 show and compare TEM micrographs obtained from the secretory regions of the normal and hypertrophied salivary glands. The analysis by light microscopy and the TEM examination revealed that the secretory portion of the normal gland is characterized by a muscular coat surrounding the gland, an epithelium that consists of a single layer of cells, and a central lumen (Figs. 3A,B). TEM observations of cross sections of this region, showed that the external muscle tissue is composed of large muscle cells, each showing numerous myofibrils (Fig. 3C). The secretory cells are separated from the muscle by a basement membrane and contain an extensive rough endoplasmic reticulum, many Golgi complexes, numerous mitochondria, some of which have electron dense granules, and a large number of secretory granules (Figs. 3D,F,G). Especially in the apical area of the epithelium, adjacent cells are in contact along their lateral plasma membranes by septate desmosomes (Fig. 3E). In addition, the secretory cells show microvilli extending in the lumen and involved in the release of secretion (Figs. 3E,H). The lumen contains an electron dense matrix, where there are numerous electron-opaque filaments (Fig. 3E). The analysis of the same region of the hypertrophied glands, infected by SGH virus, highlights that the muscular coat, surrounding the gland, undergoes a profound change in its organization (Figs. 4A,B,C). TEM micrographs show the presence of vacuoles in the muscle cells, even in the region in contact with the underlying basement membrane (Fig. 4C). A careful TEM observation reveals that the basement membrane clearly becomes an irregular and not well defined structure because of the remarkable degree of hypertrophy. In addition, it appears stratified and thicker than the same region of the healthy salivary glands and shows evident changes in the relationship with the muscle cells (Fig. 4C). Moreover, TEM investigations of the secretory region reveal a strong alteration of the cytology of epithelial cells. The glandular epithelium shows an evident vacuolization (Figs. 4A,B), cell junctions are no longer visible and the gland enlargement is caused by cellular proliferation of the secretory cells (Fig. 4D), resulting in an abnormal multilayered epithelium and a reduced gland lumen (Fig. 4A). Also, the proliferating cells show altered relationships with the degenerated basement membrane, resulting in loss of contact with it in some areas, because of the vacuolization of the cytoplasm (Fig. 4C). Numerous virus particles are scattered in the nuclei and in the cytoplasm of secretory cells (Figs. 4E,F). The figure 4E shows the morphological differences of the virus in the nucleus and in the cytoplasm; in fact, in the nucleus the virus replicates and assembles its nucleocapsid and with the transition into the cytoplasm the viral particles further develop. High magnification micrograph shows longitudinal and cross sections of the SGH virus particles in the cytoplasm of the secretory cells (Fig. 4G). In the normal salivary glands, with the transition from the secretory region to the reabsorptive region, and then, to the proximal region the diameter of the gland decreases (Fig.2A). The reabsorptive portion is localized in the thorax of the insect and at the electron microscope level Page 179

this region is characterized by an external basal lamina with a smooth outside surface and an underlying epithelial cells showing an infolded basal membrane (Figs. 5A,B). This region is very rich in microvilli that extend from epithelium into the lumen and the reabsorptive cells are involved in the reabsorption of the lumen secretion (Figs. 5A,B). Furthermore, the lumen of this portion of the gland shows a secretion less concentrated of electron-dense filaments than those of the secretory region (Fig. 5A). Finally, the morphology of the proximal region is characterized by a thin basement membrane that surround an epithelial layer. The epithelial cells are separated from the lumen by the cuticle, consisting of a lighter layer (endocuticle) and a dense dark thin layer (epicuticle) (Fig. 5C). The viral infection also affects the reabsorptive region, where the ultrastructural analysis shows a profound change of the internal organization of tissues. The TEM observations of this region revealed that the epithelium appears stratified, vacuolated and it is difficult to distinguish the boundary between epithelium and lumen (Fig. 6A). The reabsorptive cells have enormous cytoplasmic vacuoles and the microvilli seem to be destroyed or damaged (Fig. 6A). In the reduced glandular lumen it was possible to identify viral particles (Fig. 6B). The basement membrane is thicker and less compact than those of the same region of healthy salivary glands (Fig. 6C). Moreover, in the reabsorptive cells, free ribosomes, scattered strands of rough endoplasmic reticulum, and numerous degenerated mitochondria are present (Figs. 6C,D). The viral infection also affects the proximal region (Fig. 2A), but duct epithelial cells of the salivary glands undergo a minor change and alteration compared to the secretory and reabsorptive cells (Fig. 6E). In fact, no viral particles were observed in this portion of the gland. In addition, TEM observations revealed the absence of secretion in the lumen (Fig. 6E).

4. Discussion Insect salivary glands can be divided into two groups on the basis of their anatomy (House and Ginsborg, 1985). Salivary glands of the higher Diptera consist of simple tubules joined to form a common duct, whereas in other orders of insects the salivary glands are more complex, having a racemose appearance with ducts and acini. Several ultrastructural studies of dipteran salivary glands have been published (House and Ginsborg, 1985; Janzen and Wright, 1971; Martoja and Ballan-Dufrançais, 1984; Wright, 1969). In this study, it was possible to obtain a detailed reconstruction of the general organization of the salivary glands of G. pallidipes , by both SEM and TEM analyses. Along the length of this organ, we could distinguish three regions, which differ in their morphology and ultrastructure: a secretory region, a reabsorptive region, and a proximal region. A similar division of the salivary glands was observed in other insects, as in Calliphora , where the ultrastructural and physiological observations showed a secretory distal region and a proximal reabsorptive tract, along the glands (Oschman and Berridge, 1970). SEM and TEM investigation of the secretory region of the G. pallidipes salivary glands allowed an accurate description of the morphology and ultrastructure of a thick muscle tissue sheath, composed of longitudinal muscular fibers, that cover this region of glands. This muscle layer may be responsible for the contraction necessary to expel the saliva, as described in other insects (Reis et al., 2003). The presence of external muscle tissue in the salivary glands of insects is quite different. In fact, salivary glands of Anopheles stephensi (Wright, 1969), Aedes aegypti (Janzen and Wright, Page 180

1971), and of Calliphora (Oschman and Berridge, 1970) show no muscle covering. The same observation was performed in the salivary glands of Ceratitis capitata , where, in the adult glands, it has been suggested that the actin filament network may generate peristaltic contractions that move the saliva along the lumen (Riparbelli et al., 1994). Nevertheless, in the salivary glands of other insects the presence of muscle tissue has been observed, as in Cimex hemipterus (Serrão et al., 2008) and in Triatoma infestans (Reis et al., 2003). In previous studies on tsetse flies, very little attention was paid to the involvement of the muscle sheath in the secretion of the salivary glands and how the secretion is discharged from the extracellular cavity. No one has focused on the normal secretory process of the salivary glands of tsetse even though basic information about salivary gland secretion has been done on a model dipteran (Berridge and Patel, 1968; Hansen Bay, 1978; House and Ginsborg, 1985). Van Den Abbeele et al. (2010, 2007) looked at the molecular aspects of the sialome of tsetse to determine the constituents of saliva, especially in relation to possible changes in the composition of the secretion caused by infection of the trypanosome, but the overall physiology appears to have been ignored. The secretory cells of the salivary glands of G. pallidipes contain a high number of large mitochondria that have electron dense granules, characteristic of secretory cells. These cell types possess an extensive rough endoplasmic reticulum, Golgi complexes and a large number of secretory granules, suggesting an important role of these cells in the secretory process of the enzymes, ions and water, as described in previous studies (House and Ginsborg, 1985; Kokwaro et al., 1991). Studies performed on Calliphora suggest that presumably the secretory granules contain amylase, the only enzyme that has been found in the saliva of this insect. These secretion granules are probably formed in the Golgi complex, a process that has been clearly documented in other studies (Oschman and Berridge, 1970). The reabsorptive region of the salivary glands is devoid of a muscular layer and the basement membrane is in direct contact with hemolymph, similar to the reabsorptive portion of the salivary glands of Calliphora (Oschman and Berridge, 1970). The basal domain of the plasma membrane of the epithelial cells is infolded, a large number of microvilli extend from the epithelium into the lumen and the secretion in the lumen appears more dilute than that of the secretory region, suggesting an active reabsorption in this region of the salivary glands and a change in the composition of the saliva. Morphological and physiological studies performed in Calliphora support this hypothesis; epithelial cells of this region of gland are involved in the reabsorption of ions, modifying the luminal fluid, that result in a hypo-osmotic final saliva (Oschman and Berridge, 1970; Rotte et al., 2008). The proximal duct of the salivary glands was poorly investigated in tsetse flies (Kokwaro et al., 1991) and in other insects. Our results have shown that epithelial cells produce a cuticle whose morphology resembles that described in the crop of Phormia regina , an important organ of the alimentary tract of the insects (Stoffolano et al., 2010) and indicative of foregut organization. Important results of this study were obtained from the analysis of the profound morphological and ultrastructural changes of the G. pallidipes salivary glands, caused by the SGH virus. The viral infection produces a significant hypertrophy throughout the entire gland, but the highest degree of this hypertrophy affects the secretory region. Our SEM and TEM observations of this region show severe alterations of the muscular coat and muscle cells, although viral particles were not detected in these cells. The longitudinal muscle fibers lose their organization, Page 181

suggesting a degeneration of the myofibrils, which in normal muscle structure remains intact for proper function. This alteration also affects the underlying basement membrane. In fact, this structure is irregular, appears striated and, the changes of the relationships with muscle cells and proliferating epithelial cell, suggest a degeneration of cytoskeletal elements that put in communication the basement membrane with the surrounding cells. Based on these results, we can hypothesize that the contractile capacity of muscle sheath that covers the gland is greatly compromised, which should lead to a decreased ability to expel saliva. This hypothesis is further supported by the reduction of the volume of the glandular secretion in the infected salivary glands. The disease pathology also affects the reabsorptive region of the salivary glands, where the epithelial cells are rich in degenerated mitochondria, free ribosomes and some strands show interruption of the rough endoplasmic reticulum, denoting a possible altered cellular activity. Moreover, our ultrastructure observations showed the lack of microvilli in this region suggesting a compromised reabsorptive function of the salivary glands. Thus, although it is not known whether the enzyme composition and/or functions of hypertrophied salivary glands are negatively affected by the disease pathology, the results obtained in this study highlight the morphological and ultrastructural changes of the whole salivary gland, which are viral-induced, and could be related to an overall alteration of gland function. These comments are essential to correlate the action of the SGH virus in case of fly infection with the Trypanosome spp. In the normal parasite infection, trypanosomes development takes place in the salivary glands of the tsetse flies . In particular, four sequential stages of development can be recognized in the gland: (1) the uncoated epimastigote trypanosomes (with a prenuclear kinetoplast) attached by elaborate flagellar outgrowths that ramify between host microvilli; (2) uncoated premetacyclic trypomastigotes (with postnuclear kinetoplast) and reduced flagellar outgrowths; (3) nascent metacyclic trypomastigotes that have acquired the surface coat and lost their flagellar outgrowths, but remain attached to microvilli; (4) unattached coated metacyclics lying free in the saliva ready for discharge when the fly bites (Tetley and Vickerman, 1985). Previous studies reported that the impact of SGHV infection on trypanosome transmission is unclear (Burtt, 1945; Otieno et al., 1980; Whitnall, 1934). Kokwaro et al. (1991) examined the mixed infection of tsetse salivary glands, evidencing a severe cellular disintegration and a significant degeneration of trypanosomes in these cells. On the basis of the results obtained in this study, we can hypothesize that the ultrastructural alterations of the glandular epithelium, caused by viral infection, could impair the integrity of the structures necessary to the development and survival of the trypanosome in the salivary glands. Therefore, this observation may in part explain the degeneration of trypanosomes, as described above, and we can assume that the SGH virus could affect the normal ability of viral infected tsetse flies in the transmission of the trypanosome. In conclusion, the morphological and ultrastructural results provided in this study permit us to suggest the possible functional changes in the salivary glands of G. pallidipes , infected by SGH virus, as it relates to transmission of trypanosomes. Nevertheless, further studies are necessary to elucidate the mode of action of this virus, especially in relation to the possible physiological changes of the salivary glands, in an attempt to better understand how to best manage the SGH virus in tsetse flies, important vectors of Page 182

trypanosomes and to explore alternative ways to eliminate the virus from the colonies being maintained for sterile insect control and to massive release programs.

Acknowledgements We gratefully acknowledge the financial support for this study provided by the research contract N° 16419 with International Atomic Energy Agency (IAEA).

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Lietze, V.U., Abd-Alla, A.M.M., Vreysen, M.J.B., Geden, C.J., Boucias, D.G., 2011. Salivary Gland Hypertrophy Viruses: A Novel Group of Insect Pathogenic Viruses. Annu. Rev. Entomol. 56, 63-80. Martoja, R., Ballan-Dufrançais, C., 1984. The ultrastruscture of the digestive and excretory organs, in: King, R. C., Akai, H. (Eds.), Insect ultrastructure. Plenum Press, New York, vol.2, pp. 199- 261. Oschman, J.L., Berridge, M.J., 1970. Structural and functional aspects of salivary fluid secretion in Calliphora . Tissue Cell 2 (2), 281-310. Otieno, L.H., Kokwaro, E.D., Chimtawi, M., Onyango, P., 1980. Prevalence of enlarged salivary glands in wild populations of Glossina pallidipes in Kenya, with a note on the ultrastructure of the affected organ. J. Invertebr. Pathol. 36, 113-118. Reis, M.M., Meirelles, M.S., Soares, M.J., 2003. Fine structure of the salivary glands of Triatoma infestans (Hemiptera: Reduviidae). Tissue Cell 35, 393-400. Riparbelli, M.G., Callaini, G., Dallai, R., 1994. Cytoskeleton of larval and adult salivary glands of the dipteran Ceratitis capitata . Implication of microfilaments and microtubules in saliva discharge. Boll. Zool. 61, 9-17. Rotte, C., Walz, B., Baumann, O., 2008. Morphological and functional characterization of the thoracic portion of blowfly salivary glands. Arthropod Struct. Develop. 37, 372-382. Sang, R.C., Jura, W.G., Otieno, L.H., Mwangi, R.W., Ogaja. P., 1999. The effects of a tsetse DNA virus infection on the functions of the male accessory reproductive gland in the host fly Glossina morsitans centralis (Diptera; Glossinidae). Curr. Microbiol. 38, 349-354. Sang, R.C., Jura, W.G., Otieno, L.H., Mwangi, R.W., 1998. The effects of a DNA virus infection on the reproductive potential of female tsetse flies, Glossina morsitans centralis and Glossina morsitans morsitans (Diptera: Glossinidae). Mem. Inst. Oswaldo Cruz. 93, 861-864. Serrão, J.E., Castrillon, M.I., Santos-Mallet, J.R., Zanuncio, J.C., Gonçalves, T.C., 2008. Ultrastructure of the salivary glands in Cimex hemipterus (Hemiptera: Cimicidae). J Med Entomol. 45(6), 991-999. Stoffolano, J.G. Jr, Guerra, L., Carcupino, M., Gambellini, G., Fausto, A.M., 2010. The diverticulated crop of adult Phormia regina . Arthropod Struct. Develop. 39(4), 251-260. Tetley, L., Vickerman, K., 1985. Differentiation in Trypanosoma brucei : host-parasite cell junctions and their persistence during acquisition of the variable antigen coat. J. Cell Sci. 74, 1-19. Van Den Abbeele, J., Caljon, G., De Ridder, K., De Baetselier, P., Coosemans, M., 2010. Trypanosoma brucei modifies the tsetse salivary composition, altering the fly feeding behavior that favors parasite transmission. PLoS Pathog. 6(6), e1000926 . Van Den Abbeele, J., Caljon, G., Dierick, J.F., Moens, L., De Ridder, K., Coosemans, M., 2007. The Glossina morsitans tsetse fly saliva: General characteristics and identification of novel salivary proteins. Insect Biochem. Molec. 37, 1075–1085. 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. Parasitol. 118, 469-478. Whitnall, A.B.M., 1934. The trypanosome infections of Glossina pallidipes in the Umfolosi game reserve, Zuhuland. Onderstepoort J. Vet. Sci. Anim. Ind. 11, 7-21. Wright, K.A., 1969. The anatomy of salivary glands of Anopheles stephensi Liston. Can. J. Zool. 47, 579-587. Page 184

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Figure captions Fig. 1. Light and SEM microscopy of the general organization of healthy salivary glands in Glossina pallidipes. A) The salivary glands are characterized by three distinct regions: secretory, reabsorptive and proximal. Bar= 100 µm. B+C) Light microscopy magnifications of the proximal and reabsorptive regions. Bars= 20 µm. D+E) SEM of the secretory region of the salivary glands. Micrographs show the longitudinal muscle fibers surrounding this region of the salivary glands, whereas the reabsorptive and the proximal regions are devoid of the muscle fibers. Bars= 50 µm, 10 µm.

Fig. 2. Light and SEM microscopy of the hypertrophied salivary glands, infected by SGH virus, in G. pallidipes . A) The hypertrophy involves the entire gland, especially the secretory region, which the gland is entangled with tracheae, in the abdominal portion of the fly. Bar= 200 µm. B+C) Light microscopy magnifications of the junction between the two glands and the transition between the proximal and the reabsorptive regions. Bars= 20 µm. D+E) SEM of the secretory region of the salivary glands. The muscle tissue undergoes a profound alteration (i.e., muscle fibers lose their organization and are often interrupted or destroyed producing holes in the sheath). Bars=100 µm, 25µm.

Fig. 3. Light and transmission electron micrographs of the secretory region in the normal, uninfected salivary glands. A+B) Light and TEM micrographs showing the muscular sheath that surround the glands, an epithelium that consists of a single layer of cells, and a central lumen. Bars= 10µm, 1µm. C) The muscular sheath shows the muscle fibers and it is separated from epithelium by the basement membrane. Bar= 250nm. D) Magnification of the secretory cell showing the rough endoplasmic reticulum. Bar= 200nm. E) This TEM micrograph of the secretory cell provides evidence of the presence of microvilli that extend into the lumen, a septate desmosome (*) in the apical membrane of cell and a nucleus. The lumen contains an electron dense matrix, where there are numerous electron-opaque filaments surrounded by clear zones. Bars= 500nm, (*) 50nm. F+G) The secretory cell contains Golgi complexes, mitochondria, and secretory granules. Bars= 50nm, 100nm. H) Magnification of microvilli involved in the luminal secretion and cross sections of the electron-opaque filaments surrounded by clear zones. Bar= 100nm. Bm, basement membrane; Ep, epithelium; Gc, golgi complexes; Lu, lumen; M, mitochondria; Mc, muscular coat; Mv, microvilli; N, nucleus; Rer, rough endoplasmic reticulum; Sg, secretory granules.

Fig.4. Light and transmission electron micrographs of the secretory region in the hypertrophied salivary glands, infected by SGH virus. A+B) Light and TEM micrographs showing the alteration of the muscular sheath, the vacuolization of the epithelium and the significantly reduced lumen. Bars= 20 µm, 2 µm . C) In the muscle, the arrangement of muscle fibers undergoes a profound alteration and the basement membrane is degenerated. Bar= 1µm. D) This magnification of epithelium shows the cell proliferation induced by the viral infection. Bar= 5µm. E+F) The viral particles replicate and take their nucleocapsids in the nucleus and then Page 186

undergo a further development in the cytoplasm of the secretory cells. Bars= 200nm, 100nm. G) High magnification of the viral particles in the cytoplasm. Bar= 100nm . Bm, basement membrane; Ep, epithelium; Lu, lumen; Mc, muscular coat; N, nucleus; Vp, viral particles.

Fig. 5. TEM of the reabsorptive and the proximal regions of the healthy salivary glands. A+B) The reabsorptive region is bordered by a basement membrane, which is in direct contact with the hemolymph. This region shows a large number of microvilli that extend from the epithelium into the lumen and are involved in the reabsorption of the luminal secretion. Bars= 1µm, 500nm. C) The proximal region is characterized by a thin basement membrane, an epithelial layer and the epithelial cells are separated from the lumen by a cuticle that consists of an internal endocuticle and an external epicuticle. Bar= 1µm. Bm, basement membrane; Enc, endocuticle; Ep, epithelium; Epc, epicuticle; Lu, lumen; Mv, microvilli.

Fig.6. Transmission electron micrographs of the reabsorptive and proximal regions in hypertrophied salivary glands. A) The reabsorptive region shows a degenerated ultrastructure; the epithelium is vacuolated and the microvilli appear destroyed. Bar= 2µm. B) In the reduced gland lumen some viral particles are present. Bar= 250nm. C) The basement membrane is thicker and more stratified than the same region of the healthy salivary glands. Bar= 200nm. D) The reabsorptive cells are altered and show degenerated mitochondria. The arrow indicates scattered strands of rough endoplasmic reticulum. Bar= 100nm. E) A lesser degree of ultrastructural alteration seems to affect the proximal region of the salivary glands, where no viral particles were observed. Bar= 500nm. Bm, basement membrane; Enc, endocuticle; Ep, epithelium; Epc, epicuticle; Lu, lumen; M, mitochondria; Mv, microvilli; Vp, viral particles. Page 187

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Montpellier 2 University, Montpellier, France: Max Bergoin and François Cousserans

Main Collaborators:; Adly Abd-Alla,, Andrew Parker, Alan Robinson, Just Vlak, Monique van Oers, Henry Karitihi, Drion Boucias; Johannes Jehle……… at that time)….

Development of a PCR diagnostic method for the virus of salivary gland hypertrophy symptom of Glossina palldipes (GpSGHV )

Spring of 2002: first contact with Alan Robinson, Head of the Laboratory of Entomology at Seibersdorf where a tsetse colony from Ethiopia succesfully established in the late 90’s was facing high mortality associated with the salivary gland hypertrophy symptom (SGH).

June 2002: expert mission at the Seibersdorf Laboratory. An unformal collaboration is started between the Seibersdorf and the Montpellier laboratoriess to characterize the virus known to be responsible for SGH. The virus wa spurified by sucrose density gradient, the viral DNA extracted and a partial EcoRI DNA Library was produced. Among a hundred of clones, approximatly 12 kbp of DNA were sequenced and one of the sequence showing a strong homology with the P74 baculovirus envelope gene was used to design specific sets of primers to establish a sensitive diagnostic by PCR.

November 2002 : François Cousserans brougth the primers to Seibersdorf, initiated the first screening of flies and validated the PCR test. This method immediatly replaced the labour intensive and time consuming standard method of dissection and subsequent examination of salivary glands for hypertrophy. Furthermore, its high sensitivity allowed to detect the virus in asymptomatic flies.

Optimization of the PCR conditions A method of diagnostic based on the extraction of viral DNA from a single middle leg demonstrated that 100% of flies of the Uganda strain of the Entomology Laboratory at Seibersdorf were infected, most of them asymptomatically. (Abd-Alla et al. 2007)

Development of a Quantitative PCR method to analyze the virus loads in tsteste laboratory colonies Quantitative PCR analyses revealed a clear correlation between the virus copy number in pupae and their mothers. Furthermore, all the females with high copy number exhibited typical SGH, indicating the close relationship between viral loads and symptoms. (Abd-Alla et al. 2009). Demonstration that the virus excreted by saliva during blood meal was the main source of horizontal transmission of the virus (Abd- Alla et al 2010

Assembly and annotation of the GpSGHV genome (Uganda strain) A complete sequence (190,032 bp) of the GpSGHV was assembled and 160 non overlapping ORFs were identified . The annotation of the genome revealed that only 37 of these ORFs had weak homologies to genes from other invertebrate viruses, making this virus unique among large double-stranded circular DNA viruses (Abd Alla et al. 2008)

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Assembly and annotation of the GpSGHV genome (Ethiopia strain) Demonstration that the Uganda and Ethiopia viruses are strains of the same virus (to be published) Comparison of GpSGHV and MdSGHV genome sequences and organizations Demonstration that the GpSGHV and the virus of Musca domestica salivary gland hypertrophy MdSGHV) are phylogenetically related and form a distinct new family of viruses. (Garcia-Maruniak et al.2009, Abd-Alla, Boucias, Bergoin 2010)

Proposal for the classifification of MdSGHV and GpSGHV into the new family Hytrosaviriadae (Abd-Alla et al 2009b)

Production of rabbit antisera against immunodominant GpSGHV envelope proteins Proteins from ORFs 10, 47 and 96 produced in Wageningen were used to immunize rabbits. High titer antisera were produced and used in proteomic analysis of the virus (Kariithi et al 2010 and manuscript in preparation) and in sero-neutraliztion assays by feeding flies with virions incubated with the antisera (Seibersdorf)

Participation to the elucidation of 3D structure of GpSGHV virion by cryo-EM . In collaboration with Igor Orlov and Danièle Spenher at IGBMC (Strasbourg) a 3D reconstruction of GpSGHV virion is in process

References (11) Abd-Alla, A., Bossin, H., Cousserans, F., Parker, A., Bergoin, M., Robinson, A., 2007. Development of a non-destructive PCR method for detection of the salivary gland hypertrophy virus (SGHV) in tsetse flies. J. Virol. Methods 139 , 143-149.

Abd-Alla, A.M.M., Cousserans, F., Parker, A.G., Jehle, J.A., Parker, N.J., Vlak, J.M., Robinson, A.S., Bergoin, M., 2008. Genome analysis of a Glossina pallidipes salivary gland hypertrophy virus (GpSGHV) reveals a novel large double-stranded circular DNA virus. J. Virol. 82 , 4595-4611.

Abd-Alla, A., Cousserans, F., Parker, A., Bergoin, M., Chiraz, J., Robinson, A., 2009a. Quantitative PCR analysis of the salivary gland hypertrophy virus (GpSGHV) in a laboratory colony of Glossina pallidipes . Virus Res. 139 , 48 -53.

Abd-Alla, A.M.M., Vlak, J.M., Bergoin, M., Maruniak, J.E., Parker, A.G., Burand, J.P., Jehle, J.A., Boucias, D.G., 2009b. Hytrosaviridae: a proposal for classification and nomenclature of a new insect virus family. Arch. Virol. 154 , 909-918.

Garcia-Maruniak, A., Abd-Alla, A.M.M., Salem, T.Z., Parker, A.G., van Oers, M.M., Maruniak, J.E., Kim, W., Burand, J.P., Cousserans, F., Robinson, A.S., Vlak, J.M., Bergoin, M., Boucias, D.G., 2009. Two viruses that cause salivary gland hypertrophy in Glossina pallidipes and Musca domestica are related and form a distinct phylogenetic clade. J. Gen. Virol. 90 , 334-346.

Abd-Alla, A.M.M., Boucias, D.G., Bergoin, M., 2010a. Hytrosaviruses: Structure and genomic properties, in: Asgari, S., Johnson, K.N. (Eds.), Insect Virology. Caister Academic Press, Norfolk, pp. 103-121.

Abd-Alla, A.M.M., Kariithi, H., Parker, A.G., Robinson, A.S., Kiflom, M., Bergoin, M., Vreysen, M.J.B., 2010b. Dynamics of the salivary gland hypertrophy virus in laboratory colonies of Glossina pallidipes (Diptera: Glossinidae). Virus Res. 150 , 103-110.

Kariithi, H.M., Ince, A.I., Boeren, S., Vervoort, J., Bergoin, M., van Oers, M.M., Abd-Alla, A., Vlak, J.M., 2010. Proteomic analysis of Glossina pallidipes Salivary Gland Hypertrophy Virus virions for immune intervention in tsetse fly colonies. J. Gen. Virol. 91 , 3065-3074.

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Abd-Alla, A.M.M., Parker, A.G., Vreysen, M.J.B., Bergoin, M., 2011a. Tsetse Salivary Gland Hypertrophy Virus: Hope or Hindrance for Tsetse Control? PLoS Negl. Trop. Dis. 5 , e1220.

The two last papers to be published in J. Invertebr. Pathol. are missing

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FINAN REPORT

CRP: No: 14227/R0 Improving SIT for Tsetse Flies through Research on their Symbionts and Pathogens

Nguya K. Maniania The international Centre of Insect Physiology and Ecology ( icipe ) P.O. Box 30772-00100 GPO Nairobi, Kenya

Overall Objective: Understanding and exploiting interactions between tsetse flies and their microbes to enhance the efficacy and implementation of tsetse programmes with an SIT component

Specific objectives: To improve tsetse suppression technology

Output. Investigate tsetse pathogen interactions to improve control strategies

Pathogens of tsetse interact with SIT in two ways, one negative and one positive. Firstly the presence of a virus can compromise the productivity of laboratory colonies and so interfere with effective mass rearing, and secondly fungal pathogens may be used as a means to suppress tsetse populations prior to the release of sterile males. Entomopathogenic fungus (EPF) whose infection occurs through the cuticle is the most appropriate pathogen for haemathophagous insects as opposed to pathogens such as bacteria, viruses and protozoa that need to be ingested to cause infection. Studies have shown that species of tsetse flies are susceptible to fungal infection in the laboratory; but only one study has shown their use in tsetse suppression in field conditions (Maniania et al., 2006). Fungal conidia are released in the field through a contamination device (Cd) that attracts flies that come into the trap where they get contaminated with spores before they return to the environment where they may contaminate other flies during mating. This approach is called autodissemination. Fundamental to this approach is the efficient horizontal transmission of the pathogen to susceptible individuals within the insect population. Moreover, fungal infection can raise behavior changes in the host, some of which may compromise horizontal transmission of the inoculum. Horizontal transmission of entomopathogenic fungi in tsetse flies is well documented in the laboratory. However, there are no data to support this in the field. The objective of this study was, therefore, to investigate the transmission of fungal conidia between fungus-infected flies and free- healthy flies under field-cage conditions.

General Achievements Specific objectives: To improve tsetse suppression technology Output: Investigate tsetse pathogen interactions to improve control strategies

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1. Horizontal transmission of fungal inoculum between adult G. pallidipes. A contamination device (Cd) inoculated with 1.5-2.0 g of dry conidia of Metarhizium anisopliae isolate ICIPE 30 mounted on biconical trap was placed inside the Calkins and Webb’s field-cage to simulate the field conditions. Ten 7-9 day-old male flies were introduced into the trap and were forced to pass through the contaminated Cd and then exit into the cage. This group of flies was named “donor”. Equal number of 3-5 day-old female flies was released in the cage and named “first line recipient”. Flies were observed for mating. Once the pair was in copula, it was removed and placed in individual tube. After copulation, male flies were removed and returned into the cage. Another set of 3-5 day-old females was introduced in the cage and allowed to mate with already mated males. This group of flies was named “second line recipient”. The procedure was the same as described earlier. In the control, fungus-free flies were used. Test-flies (female and male) then placed individually in tubes and maintained in room conditions for 25 days. Flies were fed thrice a week on rabbit. Mortality was recorded daily and dead insects transferred to Petri dishes lined with damp filter paper to allow the growth of the fungus on the surface of the insect. Another experiment was carried using female flies as “donors”. Equal number of 7-9 day-old male flies, “first line recipient”, was released in the cage and allowed to mate. Another set of 7-9 day-old male flies was introduced in the cage and allowed to mate with already mated females. All the experimental procedure remained the same as above. Free-fungus female flies were used in the control treatments. Since only few flies managed to mate in both fungus-free and fungus-treated “donor” flies and this after many hours, control flies were not subjected to mating in the field cage because of time constraint. However, they were kept in rearing cages to monitor the health status of the test-insects used in the experiments. Ten “donor” male flies were used per treatment and the experiment was repeated four times; while 20 “donor” female flies were used and the experiment was replicated five times.

The average mortality in the control flies where male flies served as “donor” was 26.3%. All the fungus-treated “donor” male flies succumbed to fungal infection within 6 days, of which 90% showed mycosis (Figure 1). Out of the 42.5% female first line “recipient” flies that were able to mate, all of them died from fungal infection (Figure 1). Only 13% female flies managed to mate with already mated males, second line “recipient”, and all of them also succumbed to fungal infection (Figure 1). All the fungus-treated “donor” female flies died as result of fungal infection, of which 94% of the flies showed sign of mycosis (Figure 2). Compared to male ‘donor” flies, the proportion of flies that mated in female “donor” flies was significantly high. Fifty-five per cent (65%) of first line male “recipient” flies that mated, 41% of them died from mycosis (Figure 2). In the second line “recipient”, 11% male flies were able to mate with fungus-treated female “donor” flies, resulting in 10% mortality of 10 flies, of which 7% were mycosed (Figure 2).

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Figure 1. Percent mortality by mycosis of male “donor” and female “recipient” G. pallidipes following exposure of male “donor” to M. anisopliae.

Figure 2. Percent mortality by mycosis of female “donor” and male “recipient” G. pallidipes following exposure of female “donor” to M. anisopliae

2. Horizontal transmission of fungal inoculum between adult G. fuscipes. The protocol for infection of flies remains the same as described above. All the fungus- treated “donor” male and female flies succumbed to fungal infection within 6.3 days in males and more than 10 days in females (Figures 3 and 4). Thirty-five (35) and 25%

Page 200 female “recipient” flies were able to mate in the first and second line, respectively, and 95 and 100% of them died from fungal infection with mycosis within 8 and 10 days post- inoculation (Figure 3). Only 25 and 10% of male fly “recipient” managed to mate with fungus-treated female “donor” in the first and second line, respectively. Although all male flies died, only 60 and 40% of them developed mycosis, first and second line “recipient”, respectively (Figure 4).

Figure 3. Percent mortality by mycosis of male “donor” and male “recipient” G. fuscipes following exposure of male “donor” to M. anisopliae

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Figure 4. Percent mortality by mycosis of female “donor” and male “recipient” G. fuscipes following exposure of female “donor” to M. anisopliae

3. Horizontal transmission of fungal inoculum between fungus-infected irradiated male and fungus-free female G. pallidipes . Ten 7-9 day-old irradiated (100 Gy in a 60 Co) male G. pallidipes flies were forced to pass through the fungus-contaminated Cd mounted on tsetse trap in a Calkins and Webb’s field cage as described earlier. They were then transferred individually into rearing cages and a 3-5 day-old healthy female flies was released in the cage and named “first line recipient”. Once the pair was in copula, male fly was removed and returned into the cage and the female placed in individual tube. The other experimental procedure was similar as described above. Mortality in the control flies where irradiated male flies served as “donor” was 30%. All (100%) the irradiated fungus-treated “donor” male G. pallidipes flies succumbed to fungal infection within 12 days with mycosis (Figure 5). There was no significant difference in the number of female flies mating with irradiated male “donor” between the control (80%) and the fungus-treated ones (82.5%); while 55 and 42.5% of females mated in the first line and second line “recipient”, respectively. Eighty-three percent and 43% of flies that mated succumbed to fungal infection in the first and second line “recipient”, respectively (Figure 5). The rate of fungal infection observed in the present study is lower compared to previous study where most of female flies that succeeded to mate died from fungal infection. There is no explanation so far for these results. Could the irradiation have an effect on fungal conidia or there was an attenuation of virulence of conidia used in the study. Further investigation is therefore required.

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Figure 5. Percent mortality of irradiated male “donor” and female “recipient” G. pallidipes flies following exposure of male “donor” to conidia of the fungus M. anisopliae

4. Horizontal transmission of fungal inoculum from puparia emerging from fungus- contaminated soil to healthy flies. In the present study we investigate whether tsetse flies emerging from fungus-treated puparia can transfer the inoculum to healthy flies. G. pallidipes puparia were buried in 50 g of sand previously mixed with 1 g of conidia of M. anisopliae isolate ICIPE 30 . In preliminary studies, 2 g of conidia were used, resulting in acute mortalities with short survival. In the control treatment sand was fungus-free. Emerging flies were sexed and transferred in different rearing cages. This group of flies was named “donor”. Equal number of flies of opposite sex was introduced in the cage. They were observed for mating and were named “first line recipient” as above. Mortality was recorded daily for 12 days and dead insects were transferred to Petri dish for mycosis as described above. Mortality in the control flies was generally high: 30, 40 and 20% in “donor”, first and second line recipients, respectively. All the female flies that emerged from fungus-treated sand died from fungal infection with mycosis within 4 days. All male “recipient” flies succeeded to mate with fungus-treated female “donor” and all died from fungal infection with presence of mycosis. These results therefore indicate that tsetse fly puparia can be infected with fungal conidia and emerging flies would be able to transfer the inoculum to healthy ones.

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5. Effects of fungal infection of behavioral changes in tsetse flies. The effects of fungal infection was investigated on blood feeding and reproduction potential of tsetse. Glossina m. morsitans was used as example for this study. Both male and female adult G.m. morsitans were contaminated with different concentrations (3.0 x 10 4, 3.0 x 10 5 and 3.0 x 10 6 conidia ml -1) of M. anisopliae ICIPE 30 using the technique described by Maniania (1994). Flies were then transferred individually into in small aerated tubes (3 x 5 mm) and offered blood meal by feeding them on rabbit at 2, 5, 7, 9 and 12 days post-infection. The blood meal size was calculated by weighing flies before and after the meal. The weight of blood meal by female flies was generally higher than that recorded for male flies. The effect of fungal infection on blood meal intake by both male and female flies was only observed at the higher concentration of 3.0 x 10 6 conidia ml -1. Fungal infection had also an effect of the number of puparia produced by females. For instance, “recipient 1” and “recipient 2” female flies that mated with fungus-treated males produced fewer pupae than did the fungus-free flies.

Constraints The main constraint in implementing the activities of the project was the supply of tsetse flies. A large number of tsetse flies was required to carry out bioassays. Since tsetse rearing is no longer icipe’s core activity, only limited number of flies could be obtained from the Mass Rearing and Quarantine Unit of icipe . I had therefore to rely on external sources which were either not reliable or too expensive. For instance, we only got three consignments of flies from Slovakia, of which two had viable puparia. Flies were also bought once from Trypanosomiasis Research Centre, KARI Muguga at US$2 per fly. Another attempt to purchase flies failed.

Conclusions We have demonstrated horizontal transmission of inoculums between flies in field-cage, simulating field conditions, a prerequisite to the success of autodissemination approach. Nonetheless numerous research challenges remain. Laboratory experiments have demonstrated the complementary action of EPF on mortality, reduction in blood feeding and reproduction potential. On the other hand, the effect of fungal infection by M. anisopliae on mating behavior was investigated in a previous study and there were no significant differences in competitiveness between fungus-treated males and fungus-free males. The mean duration of copulation was also similar (Maniania unpubl. data). It may be therefore possible to use sterile males to disseminate spores of EPF since both sterilization (Abila et al., 2003) and fungal infection does not affect mating behavior of the flies. The impact of fungal infection on trypanosome development and transmission is an area for future research. This has been shown in malaria parasite in (Blanford et al., 2005; Scholte et al., 2005).

Acknowledgments

I would like to thank Mr. J.O. Opere for technical assistance, Dr. P. Takac, Institute of Zoology, Slovakia, for providing flies, the Kenya Agricultural Research Institute/Trypanosomiasis Centre (KARI/TRC), Muguga, for irradiation of flies, and IAEA for financial support.

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Final Report: The role of pathogens and symbionts of Glossina brevipalpis and G. austeni in South Africa

Otto Koekemoer, March 2012

Abstract

Field samples of G. brevipalpis and G. austeni were collected from areas covering the total known distribution of the two species in South Africa and from the Mlawula National park in Swaziland. DNA extractions were made from whole flies and used to determine the presence of both Wolbachia and GpSGHV by PCR. The GpSGHV-specific PCR revealed only a single infection in colonized flies and very low levels of infection in wild specimens. G. austeni had an average Wolbachia infection rate of 95% while G. brevipalpis only showed infection in 61%. Typing of the symbiont from Wolbachia positive flies using the MLST approach revealed the presence of at least two genotypes in G. brevipalpis , one of which has a new sequence on the hcpA locus. Further genetic distinctions could be made between Wolbachia infecting G. brevipalpis and G. austeni from different locations using a high resolution melting profile analysis of wsp -gene PCR products.

1. Introduction South Africa has historically been inhabited by four species of tsetse. These included G. morsitans morsitans, G. pallidipes, G. austeni and G. brevipalpis . A combination of the rhinderpest outbreak in the 1890’s, game and forest clearing for farming purposes and insecticide spraying led to the eradication of two species along the northern and eastern borders of the country. Today only G. austeni and G. brevipalpis remain in an area in the north eastern part of KwaZulu-Natal (Kappmeier et al., 1998). The area infested is discontinuous but estimated at approximately 18 000 km 2 and is compromised of communal and commercial farming operations surrounding a number of private and national game parks and protected areas. Starting in 1990 and since the farming areas that surround the game parks have undergone several outbreaks of animal trypanosomiasis (nagana) affecting cattle. With this in mind integrated approaches are sought to suppress or eradicate tsetse populations involved in the transmission of the disease. One strategy that has been identified as part of such an approach is to use SIT to suppress populations to be controlled in combination with other methods like trapping or areal spraying. In support of these efforts, studies on the natural symbionts and pathogens of tsetse in South Africa have been undertaken.

The aim of this project was to collect flies from different locations, and to test these for the presence of pathogens similar to, or the same as the virus that has been identified in G. pallidipes (Odindo et al., 1986). It is known that both G. brevipalpis and G. austeni are infected

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with Wolbachia but the levels of infection are not known, neither are the strains involved. Molecular methods are available by which the presence of both viral and bacterial ( Wolbachia ) DNA can be indicated (Abd-Alla et al., 2011, Werren and Windsor, 2000). For the genetic characterization of the Wolbachia strains that are present in South African tsetse, the MLST protocol of Balso et al. (2006) and a PCR-HRM assay of Henri and Mouton (2011) were used. This report presents the findings up to date.

2. Methods and Results 2.1. Field collections H-traps (Kappmeier, 2000) were set up in areas that were previously surveyd in in KwaZulu-Natal (Kappmeier et al., 1998) and the Mlawula National Park in Swaziland and were known to be inhabited by tsetse. Traps were baited with a 1 : 8 ratio of 1-octen-3-ol and 4-methyl phenol and acetone and catches were collected daily. The flies were either used directly for dissection or stored in 96% EtOH until further processing took place. Multiple traps (4 – 10 set up at distances of between 200 – 400 m apart) were placed at the locations indicated in Figure 1.

2.2. DNA extraction Flies caught in H-traps were collected in 96% EtOH and sorted at the field station. Specimens were placed in fresh 96% EtOH and kept until use. Individual flies were dried on clean tissue paper and rehydrated in normal saline. The bodies without wings and heads or the carcasses remaining after dissection, were ground using a micropestle in 300 – 500 µL sterile PBS. 200 µL of this suspension was used for DNA extraction using the Roche High Pure PCR Template Preparation Kit as per included instructions. The DNA was suspended in 300 µL elution buffer and kept at -20 °C.

2.3. Microscopic examination of salivary glands Flies were dissected in PBS using instruments that were cleaned between each individual dissection. The flies were examined for salivary gland hypertrophy and reproductive tissues were removed from some of the specimens and placed in 100% EtOH for work on symbiont characterization. None of the flies (both field and colony flies) showed any signs of salivary gland hypertrophy. In less than 10 cases where colonized flies were examined, salivary glands with an abnormal, opaque appearance was observed (Fig. 2). It is unknown if this is related to any infection. A total of 185 wild flies and 203 colony flies were examined.

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Figure 1 . Areas where H-traps were placed in north-eastern KwaZulu-Natal and Swaziland from where tsetse were collected during the study.

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Fig. 2. A salivary gland with an unusual opaque appearance from a colonized G. brevipalpis .

2.4. SGHV detection by PCR The initial primers that were used as described by Abd-Alla et al. (2007) gave indications of a very high rate of SGHV infection in field flies. Some of the PCR products were sequenced and it was found that the amplification products were non-specific artifacts. These results were discarded and degenerate primers (Abd-Alla et al., 2011) were used. The PCR was followed as described using the PIF1-2 and P74 primer sets. Nine % of field G. austeni and 29 % of field G. brevipalpis tested gave amplification products, but only when using the PIF primers. No positive results were obtained with the P74 primers.

Fig. 3. Agarose gel electrophoresis of amplification products after GpSGHV-specific PCR using the Pif1-2 primer set. Arrows indicate the positions of groups of putative positives.

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These amplification products appeared very weak when compared to the positive controls on agarose gel electrophoresis, but were in the correct size range (Fig. 3).

2.5. Wolbachia detection and characterization 2.5.1. Species-specific PCR Wolbachia genus specific primers W-Specf (CATACCTATTCGA-AGGGATAG) and W-Specr (AGCTTCGAGTGAAACCAATTC) that amplify a 438 bp fragment of the 16S rDNA gene (Werren & Winsor, 2000) were used to detect the presence of Wolbachia DNA in the genomic DNA extracts. A set of general eukaryotic 28S rRNA primers were used as a control for the quality of the extracted DNA extraction. Only samples that were positive for this product were included in the calculation of the results. The prevalence are shown per site in Table 1. On average the rate of Wolbachia infection are 95% and 61% for G. austeni and G. brevipalpis respectively.

Table 1. Results of a survey for the presence of Wolbachia in wild tsetse using the PCR test described by Werren and Windsor (2000).

Location % Pos. Location % Pos.

G. austeni G. brevipalpis Mkuze 100 Hellsgate 83 Muzi Swamp 83 HI South 56 Phinda 90 HI North 63 Mbazwane 90 False Bay Park 75 False Bay Park 100 Kosi Bay 100 Tembe 100 Tembe 75 Charter’s Creek 100 St Lucia 55 Swaziland 100 Ndumu 0 Charter’s Creek 44

2.5.2. Wolbachia genotyping Typing by MLST DNA extracts from flies that were shown to be Wolbachia positive were selected for typing of the symbiont by MLST according to the methods described by Baldo et al., 2006.

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Fig. 4. MLST amplification products grouped per individual fly. Five housekeeping genes and the 28S rRNA-gene were partially amplified for each individual. The 5 th position in each case (fpbA) shows no amplification. The last lane in each case is the 28S rRNA-gene control.

The 28S rRNA-gene primers were again included as internal control for quality of the DNA preparations. In most cases the control PCR product and products from the five housekeeping genes were amplified (Fig. 4). There was a consistent problem with the fbpA locus not amplifying with any of the samples. In some instances only some of the loci amplified from a specific individual. The fbpA-primers were changed for the set described to work with super group B ( http://pubmlst.org/wolbachia/info/amp_seq_single.shtml ) and this locus could be amplified for sequencing. The PCR products were directly sequenced in both directions. Sequence data were aligned and compared with data available in the Wolbachia MLST databases. Forty loci were sequenced from eight individual tsetse. The results of the typing by sequencing are summarized in Table 2.

wsp-gene analysis by HRM Another method of identifying genetic variation in the genomes of Wolbachia symbionts was investigated. The method makes use of sequence variation in hyper variable areas of the wsp-gene of Wolbachia (Henri and Mouton, 2012) and is based on of a set of primers

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Table 2. MLST per fly per location. Allele profile numbers are as reported by the Wolbachia MLST website query Locus gatB CoxA hcpA ftsZ fbpA G.b. Colony 128 109 - 98 20 G.a. Colony 128 New (109) - 98 20 G.b. Hluhluwe South x2 14 129 New (23) 56 15 G.b. Hluhluwe North - - 127 98 20 G.a. Mkuze 128 109 127 98 20 G.a. Charter’s Creek 128 109 127 98 20 G.a. Swaziland 128 109 127 98 20 (-) Sequences that were not full length new Indicates an allele that is not presented in the MLST database. The number is that of the closets allele.

that amplify a 610 bp region of the wsp-gene. A dilution of this PCR product is then used to carry out a nested PCR with primers that amplify smaller parts of the hyper variable regions (HVR) of the gene. These nested products are used in the high resolution melting analysis in the presence of a saturating fluorescent dye in a real-time PCR format. Pre- amplification was assessed on an agarose gel and positive samples were used for the nested PCR and HRM analysis.

The most obvious distinction in melting profiles was obtained when the HVR3-region was used. The HVR2-region revealed no clear distinctions between groups of flies. With G. brevipalpis two groupings emerged: the first contained flies from the Tembe, Hellsgate, False Bay park and Charter’s creek areas while the other consisted of flies from Kosi bay, Hellsgate and Hluhluwe (Fig. 5A). In the case of G. austeni only one fly from Swaziland showed a different melting profile (Fig. 5B).

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A

B

Figure 5. HRM melting difference plots of amplification products related to HVR3-region of the wsp gene. G. brevipalpis (A) and G. austeni (B).

3. Conclusion The total area in South Africa and one known location in Swaziland as well as colony tsetse were surveyed for the presence and signs of SGHV and Wolbachia in G. austeni and G. brevipalpis using PCR methods and physical examinations.

Except for one G. brevipalpis , there was no positive results when testing tsetse from the colonies at Onderstepoort. One set of degenerate primer (pif-2) revealed results that might indicate the presence of viral DNA. This will have to be confirmed using sequencing of the PCR products as p74-based PCR revealed no positives. This might be the result of nucleotide

Page 213 variations in the sites where the PCR-primers are to bind. No physical evidence of salivary gland hypertrophy could be observed in any case where freshly collected flies were examined.

The Wolbachia surveys revealed a very high rate of infection in G. austeni . In multiple areas the infection rate was 100%. The lowest rate of infection in this species was 83%. In G. brevipalpis there was much bigger variation ranging from areas that showed a 0% infection rate to areas where 91% of the flies were positive. Selected positive individuals were used to carry out molecular typing of their Wolbachia symbionts using two different assays. In the first analysis the MLST protocol of Baldo et al., (2006) was used. Results from G. austeni were the same as recently reported by Doudoumis et al. (2012). No variation was observed between different origins, no new alleles could be detected except for one seemingly new allele for the CoxA locus of a colony fly. G. brevipalpis , however, showed variation in one set of samples from the southern part of the Hluhluwe Imfolozi Park. Sequence variation was observed on all 5 loci between Wolbachia alleles from these flies and specimens from other locations in South Africa. Furthermore, flies from this location showed a new allele, not currently present in the MLST database on the hcpA gene.

A paper by Henri and Mouton (2011) describes a methods that makes use of PCR and HRM to rapidly analyse Wolbachia for genetic diversity. It does not rely on sequencing and targets the highly variable wsp-gene. One of the regions on this gene, HVR3, showed genetic diversity within the Wolbachia populations that were tested, and revealed variation between flies that were not seen using the MLST approach. In the case of G. brevipalpis a distinction could be seen between two groups of flies, one containing specimens from the Kosi bay, Hellsgate and Hluhluwe areas, and the other specimens from Tembe, Hellgate, False Bay park and Charter’s Creek. This not only confirms the genetic diversity that was observed with the MLST analysis, but it shows that there is variation within at least one area viz. Hellsgate (see Fig. 5). Wolbachia from G. austeni from Swaziland showed no genetic variation using MLST but a clear variation was observed in one specimen from this location. Again, there was variation within the group, as other specimens from Swaziland showed the same profile as that of the Wolbachia that were from flies collected in South Africa (Fig. 5B).

In summary, although the virus-specific PCR showed some results that can be interpreted as positive for the presence of the virus, this needs to be confirmed by sequencing of the amplicons. Wolbachia infection rates were established for the different areas under survey and genetic typing has revealed genetic variation in symbionts from both host species occurring in South Africa and Swaziland.

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4. References

Abd-Alla A, Bossin H, Cousserans F, Parker A, Bergoin M, Robinson A. 2007. Development of a non-destructive PCR method for detection of the salivary gland hypertrophy virus (SGHV) in tsetse flies. J Virol Methods . 139:143-149.

Abd-Alla AM, Salem TZ, Parker AG, Wang Y, Jehle JA, Vreysen MJ, Boucias D. 2011. Universal primers for rapid detection of hytrosaviruses. J Virol Methods . 171:280-3.

Baldo L, Dunning Hotopp JC, Jolley KA, Bordenstein SR, Biber SA, Choudhury RR, Hayashi C, Maiden MC, Tettelin H, Werren JH. 2006. Multilocus sequence typing system for the endosymbiont Wolbachia pipientis. Appl Environ Microbiol . 72:7098-110.

Doudoumis V, Tsiamis G, Wamwiri F, Brelsfoard C, Alam U, Aksoy E, Dalaperas S, Abd-Alla A, Ouma J, Takac P, Aksoy S, Bourtzis K. 2012. Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina). BMC Microbiol . Jan 18;12 Suppl 1:S3

Henri H, Mouton L. 2012. High-resolution melting technology: a new tool for studying the Wolbachia endosymbiont diversity in the field. Mol Ecol Resour . 12:75-81.

Kappmeier K. 2000. A newly developed odour-baited "H trap" for the live collection of Glossina brevipalpis and Glossina austeni (Diptera: Glossinidae) in South Africa. Onderstepoort J Vet Res . 67:15-26.

Kappmeier K, Nevill EM, Bagnall RJ. 1998. Review of tsetse flies and trypanosomosis in South Africa. Onderstepoort J Vet Res . 65:195-203.

Odindo MO, Payne CC, Crook NE, Jarrett P. 1986. Properties of a novel DNA virus from the tsetse fly, Glossina pallidipes . J Gen Virol . 67:527-36.

Werren JH, Windsor DM. 2000. Wolbachia infection frequencies in insects: evidence of a global equilibrium? Proc Biol Sci . 267:1277-85.

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Final report

Institute of Zoology, Slovak Academy of Sciences. Slovakia: Takac, P.

Collaborators: Aksoy, S., Malacrida, A., Abd-Alla, A.

Title: The efect of antibiotic and yeast treatment on Glossina genus tsetse flies

Abstract During the CRP project and within the contract agreement we focused on the provision of tsetse biological material to research groups. We had been supply of 10,000 tsetse fly puparia as required to participants of the CRP D42012 "Improving SIT for tsetse flies through research on their symbionts and pathogens in 14 shipments per year. a. 200 puparia shipped from G. m. morsitans, G. pallidipes and G. f. fuscipes colonies monthly to S. Aksoy, Yale. b. Four allotments of 200 puparia from G. m. morsitans shipped to A. Malacrida, Pavia. c. Puparia (1000 for each species) from G. pallidipes and G. fuscipes shipped to N. Maniana, ICIPE. At the same time we set up the collaborative experiments with Aksoy´s lab., concerning the supplementation of the host diet with yeast , different vitamine mixtures and different antibiotics, to study the influence of this treatment to microbial flora, fecundity and mortality of G. morsitans morsitans and G. fuscipes fuscipes colonies.

Introduction Glossinidae (Diptera) are vectors of African pathogenic trypanosomes, which are of medical and economic importance. They are exclusively haematophagous and this higly restricted nutritional ecology has resulted in obligate adaptations with symbiotic bacteria in tsetse. The endosymbionts provide nutritional supplements, in the absence of which females are becoming sterile. Tsetse´s dependence on its obligate microbiota for reproduction provides a weak link in its biology, and may generate alternative control strategies that should be explored. These endosymbionts have also an important role in parasite transmission. The mutual interactions result in host immune regulation and nutritional environment in the midgut and may also contribute to the vector susceptibility traits observed in field populations. This way, the endosymbionts may be potentially an efficient target for controlling tsetse fly vectorial competence and consequently sleeping sickness. The three organisms have been already characterized for Glossina genus, Wigglesworthia glossinidia, Sodalis glossinidius and Wolbachia pipientis and all three symbionts are, in essence, materially associated with the metabolism of B-complex vitamins essential for tsetse survival, the effect of yeast based meals and different B- complex vitamins meals on fitness and fecundity was analyzed.

Results 1. Provision of biological tsetse material to CRP research groups During the CRP project and within the contract agreement we focused on the provision of tsetse biological material to research groups. We had been supply of 10,000 tsetse fly Page 216 puparia as required to participants of the CRP D42012 "Improving SIT for tsetse flies through research on their symbionts and pathogens in 14 shipments per year. a. 200 puparia shipped from G. m. morsitans, G. pallidipes and G. f. fuscipes colonies monthly to S. Aksoy, Yale. b. Four allotments of 200 puparia from G. m. morsitans shipped to A. Malacrida, Pavia. c. Puparia (1000 for each species) from G. pallidipes and G. fuscipes shipped to N. Maniana, ICIPE.

2. Impact of antibiotic, yeast and yeast vitamims mixture treatment on host fecundity and lifespan We evaluated the effect of continuous per os treatment of fertile females with the tetracycline and ampiciline based antibiotics and yeast extracts or vitamins mixture diets by measuring the total number of larvae deposited per group over the course of three deposition cycles. To determine if the tetracycline and ampiciline antibiotics have a prolonged effects on the process of metamorphosis within the larvae or pupae. The hatching rate of progeny was determined in these groups. At the same time we measured the influence of antibiotic and yeast treatment on fly lifespan. We measured the effect of tetracycline, tetracycline and yeast, ampiciline, ampiciline and yeast ; only yeast and different vitamine mixture treatment on pupae production. Under optimum conditions the first gonotrophic cycle takes about 20–22 days for development from egg to parturition. In subsequent gonotrophic cycles females produce a larva every 9 to 11 days. Ampiciline treatment does not reduce fecundity since it does not damage Wigglesworthia resident within bacteriocytes in the midgut, unlike tetracycline, which clears all bacteria including Wigglesworthia and Wolbachia and induces sterility. Accordingly, ampicillin- receiving flies remained fecund while tetracycline receiving flies were rendered sterile. Yeast extract (10% w/v) provisioning of the blood meal rescued fecundity of the females receiving tetracycline to similar levels as that of wild type and ampicillin receiving flies (65%, 55% and 64% over the first gonotrophic cycle and 53%, 58% and 49% over the second gonotrophic cycles, respectively). However, yeast provisioning at 10% w/v had a cost on fecundity when compared to flies maintained on normal blood meals, (92% versus 55% over the first gonotrophic cycle and 92% and 58% over the second gonotrophic cycle, respectively). Nevertheless, yeast supplementation was able to rescue the tetracycline-induced sterility to levels comparable to those observed for Gmm Wt receiving yeast or ampicillin supplemented blood meals, respectively.Thus yeast supplemented dietary regiment allowed us to develop two lines to analyze the functional role of Wolbachia symbionts in tsetse biology; one lacking all symbionts ( Gmm Apo ) and another lacking Wigglesworthia but still retaining Wolbachia and Sodalis (Gmm Wig−). The Gmm Apo progeny resulting from the first and second depositions of tetracycline treated mothers were tested for the presence of Sodalis, Wigglesworthia and Wolbachia by a bacterium-specific PCR-assay. The PCR-assay demonstrated the absence of all three symbionts as early as the first deposition in both the male and female Gmm Apo adults. The absence of Wolbachia from the reproductive tissues of Gmm Apo females was also verified by Fluorescent In Situ Hybridization (FISH) analysis. In contrast, Wolbachia was present in egg chambers during both early and late developmental stages in Gmm Wt females. Longevity of F1 Gmm Apo females was compared to that of Gmm Wt adults maintained on Page 217 the same yeast-supplemented blood meal over 40 days (two-gonotrophic cycles). No difference (X 2 = 0.71, df = 1, P = 0.4) was observed in survivorship comparisons between the two groups. The second line ( Gmm Wig −) generated from ampicillin treated females still retain their Wolbachia and Sodalis symbionts, while lacking both Wigglesworthia populations as evidenced by FISH and PCR amplification analysis. When maintained on yeast-supplemented blood, this line (similar to Gmm Apo ) also did not display any longevity differences from the Gmm Wt adults sustained on the same diet.

Conclusions Through understanding the mechanisms by which tsetse endosymbionts potentiate trypanosome susceptibility in tsetse, it may be possible to engineer modified endosymbionts which, when introduced into tsetse, render these insects incapable of transmitting parasites. In our study we have assayed the effect of two different antibiotics on the endosymbiotic microflora of tsetse Glossina genus. We showed that the antibiotics, ampiciline and tetracycline, have a different impact on tsetse fecundity, pupal emergence, kortality, efectively rendering these insects sterile. Using the yeast extracts and different vitamine mixtures partially eliminate the antibiotic efect. Base on our results we can consider the yeast diet as the most appropriate to sustain optimal colony of endosymbiont-free flies.

1. Paper citation:

Alam U , Medlock J , Brelsfoard C , Pais R , Lohs C , Balmand S , Carnogursky J , Heddi A , Takac P , Galvani A , Aksoy S. (2011) Wolbachia Symbiont Infections Induce Strong Cytoplasmic Incompatibility in the Tsetse Fly Glossina morsitans . PLoS Pathog 7(12): e1002415. doi:10.1371/journal.ppat.1002415

Paper link : http://www.plospathogens.org/article/info%3Adoi%2F10.1371%2Fjournal.ppat.1002415; jsessionid=554224F4111AA0B7AB53D29EC2D2BB5A

2. Paper citation: DOUDOUMIS V, TSIAMIS G, WAMWIRI F, BRELSFOARD C, ALAM U, AKSOY E, DALAPERAS S, ABD-ALLA Adly, OUMA J, TAKAC P, AKSOY S, BOURTZIS K. (2012) Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina). BMC Microbiology, vol. 12, suppl. 1, s3; 13pp. ISSN 1471-2180.

Paper link: http://www.biomedcentral.com/1471-2180/12/S1/S3 Page 218

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WORKING PAPER

Generation and evaluation of polyclonal antibodies against hytrosaviruses

Wuhan Institute of Virology, Chinese Academy of Sciences: Zhihong Hu

Collaborators: Drion G. Boucias, Just M.Vlak

Introduction

SGHV infection is responsible for the abnormal mortalities of the mass rearing host fly. PCR detection indicated that there no negative fly in the colony till now and most of the flies have the asymptomatic infection. The objectives of our research are to generate SGHV specific antibodies that can be used for immunohistological study of SGHV infection, as well as to evaluate if any antibodies could neutralize the virus oral infection.

During our study selected ORFs of MdSGHV and GpSGHV were cloned and expressed in E. coli . Antibodies against ORF22, ORF86, ORF96 and IAP from MdSGHV, and PIF1, PIF2, ORF140 and ORF261R from GpSGHV were successfully generated. Antibodies against ORF22, ORF86 and ORF96 could be used for pathology study and bioassay showed that antibodies against ORF96 could suppress the oral infection of Polyclonal antibodies from rabbit against these expressed proteins were generated. Western blot and EM were used to evaluate the antibodies.

Results

（（（1）））Antibodies against ORF22, ORF86, ORF96 and IAP from MdSGHV were successfully generated

Sequences analysis showed that orf22, orf86 and orf96 encode major structure proteins of MdSGHV. These genes as well as a putative important gene iap were chosen for expression and antibodies generation. These genes cloned into the transfer vector were provided by Dr. Drion G. Boucias. After the restriction enzyme digestion, the genes were harvest and subcloned into the prokaryotic expression vector pProExHT (Invitrogen).

Working Paper-Huzh 2012 Page 220

The correct plasmids were cultured for expression in E. coli and the proteins were purified. 400 µg purified proteins were introduced to immune each rabbit. 4 weeks later, 200 µg purified protein were boosted to the rabbit. One week later, the second boost was introduced and the rabbit sera were collected 1-2 weeks later. The sera were purified and Western blot showed that antibodies ORF22, ORF86, ORF96 and IAP from MdSGHV were successfully generated. These antibodies were to CRP collaborator Dr. Drion G. Boucias for further analyses.

（（（2）））Antibodies against PIF1, PIF2, ORF140 and ORF261R of GpSGHV were successfully generated

The plasmid with the genes chitinase, pif1 , pif2 , pif3, orf140, orf118L, orf261R, and orf415R of SGHV were obtained from Prof. Just M. Vlak. These genes were cloned in to prokaryotic expression vector pET28 (Novagen). After transfer the constructed plasmid in to expression E. coli strain BL21, only PIF1, PIF2, ORF140 and ORF261R were highly expressed, but not the Chitinase and PIF3. Using the methods mentioned above, the antibodies against PIF1, PIF2, ORF140 and ORF261R of GpSGHV were successfully generated and submitted to CRP collaborator Dr. Just M. Vlak for further analyses.

(3) Antibodies against ORF22, ORF86 and ORF96 of MdSGHV could be used for pathology study

Dr. Drion G. Boucias’s lab tested the antibodies against MdSGHV. The result showed that antibodies anti-ORF22 and anti-ORF86 could reacted with MdSGHV infected gland and purified virus by Western blots, but anti-IAP could not.

Anti-MdSGHV22, anti-MdSGHV86 and anti-MdSGHV96 antibodies were used for immunoelectron microscopy. The results showed that anti-ORF86 reacted to the nucleocapsid while anti-ORF96 reacted to the membrane of the virion (Fig. 1 and Fig. 2). Page 221

Fig. 1. The immuno-EM of MdSGHV virion by using anti-MdSGHV86 antibody (provided by Dr. Boucias).

Fig. 2. The immuno-EM of MdSGHV virion by using anti-MdSGHV96 antibody (provided by Dr. Boucias).

（（（4）））Anti-MbHV96 antibody could suppress the level of oral infections

Newly emerged house flies were treated with anti-MdSGHV MdSGHV96a, b, and c sera and the result showed that the treatment could suppress but not eliminate virus infection (Fig. 3).

Working Paper-Huzh 2012 Page 222

Fig.3. Anti-MbHV96 antibody could suppress the level of oral infections. Percent MdSGHV-infection induced by oral treatment of newly emerged house flies with virus inoculum that was either untreated or mixed with control rabbit serum or with a cocktail of anti-MdSGHV96a, b, and c sera. (provided by Dr. Boucias).

Conclusions

In summary, polyclonal antibodies against orf22 , orf86, orf96 and iap genes from MdSGHV, and pif1 , pif2, orf140 and orf261R from GpSGHV were successfully generated. Western blot and IEM confirmed that MdSGHV86 is a main nucleocapsid protein and MdSGHV96 is a main envelope protein. Bioassays demonstrated that the MdSGHV96 polyclonal antibodies significantly decreased per os infectivity of MdSGHV. Therefore, the research provides useful antibodies for pathology study of hytrosaviruses, and it proved the principle that neutralizing antibodies could be used to prevent the per os infectivity of hytrosavirusese. The objectives of the research are accomplished.

Publication:

D. G. Boucias , F. Deng, Z. Hu , A. Garcia-Maruniak, and V.-U. Lietze. 2012. Immunological Analysis of Structural Proteins from the Musca domestica Salivary Gland Hypertrophy Virus. Journal of Invertebrate Pathology. In press . Page 223

Yale University, USA: Aksoy, S. Collaborators: Heddi, A., Malacrida, A., Bourtzis, K., Wolfgang, M and Takac, P.

Abstract : During this CRP program, we worked on the role of tsetse symbionts Wigglesworthia and Wolbachia on host fecundity and reproduction and the impact of tsetse symbionts on host immunity. Although the presence of symbiotic microbes, especially Wigglesworthia was shown to be essential for fecundity, the molecular basis of its symbiotic role remained unknown. Similarly, although laboratory and natural populations of tsetse have been shown to harbour Wolbachia , the functional role of Wolbachia in tsetse biology remained unknown. We showed that loss of fecundity due to Wigglesworthia can be rescued by supplementing the host diet with yeast. We developed and used the tsetse lines that are free of Wigglesworthia and all symbionts, respectively to investigate host-symbiont interactions. Our results showed that absence of Wigglesworthia from larval development adversely impacts cellular immunity in emerging adults and makes tsetse unusually susceptible to trypanosome infections. Our results also showed that Wolbachia infections confer strong CI in tsetse and mathematical modelling support the use of CI to derive genetically modified symbionts to replace susceptible populations in nature. Finally, we investigated the density dynamics of symbionts and SGHV in lines that lack Wigglesworthia only or that are aposymbiotic. Our results show that the tsetse host strongly regulates the density dynamics of its microbiome and that absence of Wigglesworthia compromises Sodalis fitness and that SGHV is negatively impacted by host immune responses elicited against the symbionts, especially Wolbachia . We have also collaborated with other members of this CRP on, Wolbachia infection prevalence in natural tsetse populations (DOUDOUMIS et al. 2011), presence of polyandry in tsetse (BONOMI et al. 2011) and on the molecular aspects of SGHV proteome (KARIITHI et al. 2011).

Introduction: Tsetse flies have a highly regulated and defined microbial fauna made of 3 bacterial symbionts (obligate Wigglesworthia glossinidia , commensal Sodalis glossinidius and parasitic Wolbachia pipientis) in addition to a DNA virus ( Glossina pallidipes Salivary gland Hypertrophy Virus, GpSGHV). The microbiome are vertically transmitted from mother to offspring during this insect’s unique viviparous mode of reproduction. Many animals including insects such as mosquitoes rely on the presence of symbiotic bacteria for proper immune system function. However, the molecular mechanisms that underlie this phenomenon are poorly understood including their role in tsetse. We summarized the state of this knowledge in a review paper (WEISS and A KSOY 2011) Similarly, although the presence of Wolbachia has been found in tsetse, its role on tsetse reproduction is unknown. This is largely because elimination of tsetse’s symbiont Wigglesworthia renders tsetse sterile preventing the development of lines where these functions can be experimentally tested. Finally, we hypothesized that there is a strong community influence that determines the population dynamics of individual symbionts as well as SGHV. We describe below briefly the experiments performed that allowed us to investigate these interactions.

1. Development of lines that lack tsetse symbionts . We investigated the transmission route of tsetse’s endosymbionts which showed the presence of two populations of Wigglesworthia , an intracellular state in the bacteriome and an extracellular state in the milk (ATTARDO et al. 2008). This knowledge allowed us to develop lines that are cured of their Wigglesworthia infections (PAIS et al. 2008).

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2. Role of symbionts on host immunity . We characterized the immune phenotype of tsetse that develop in the absence of all of their endogenous symbiotic microbes. Larval tsetse that undergo intrauterine development in the absence of their obligate mutualist, Wigglesworthia , exhibit a compromised immune system during adulthood (WEISS et al. 2011). We found that aposymbiotic tsetse ( Gmm Apo ) present a severely compromised immune system that is characterized by the absence of phagocytic hemocytes and atypical expression of immunity-related genes. The susceptible phenotype exhibited by Gmm Apo adults can be reversed when they receive hemocytes transplanted from wild- type donor flies prior to infection (WEISS et al. 2012). Furthermore, the process of immune system development can be restored in intrauterine Gmm Apo larvae when their moms are fed a diet supplemented with Wigglesworthia cell extracts. Our finding that molecular components of Wigglesworthia exhibit immuno-stimulatory activity within tsetse is representative of a novel evolutionary adaptation that steadfastly links an obligate symbiont with it’s host.

3. Role of Wolbachia in tsetse biology . Infections with the parasitic bacterium Wolbachia are widespread in insects and cause a number of reproductive modifications, including cytoplasmic incompatibility (CI). There is growing interest in Wolbachia, as CI may be able to drive desired phenotypes such as disease resistance traits, into natural populations. Although Wolbachia infections had been reported in tsetse, their functional role was unknown. This is because attempts to cure tsetse of Wolbachia by antibiotic treatment damages the obligate mutualist Wigglesworthia , without which the flies are sterile. Using the Wolbachia free and still fertile tsetse lines, we performed mating experiments for the first time, which provides evidence of strong CI in tsetse. We have incorporated our empirical data in a mathematical model and show that Wolbachia infections can be harnessed in tsetse to drive desirable phenotypes into natural populations in few generations. This finding provides additional support for the application of genetic approaches, which aim to spread parasite resistance traits in natural populations as a novel disease control method. Alternatively, releasing Wolbachia infected males can enhance Sterile Insect applications, as this will reduce the fecundity of natural females either uninfected or carrying a different strain of Wolbachia (ALAM et al. 2011).

4. Community dynamics regulating symbiotic densities. It has been possible to rear flies in the absence of either Wigglesworthia or in totally aposymbiotic state by dietary supplementation of tsetse’s bloodmeal. In the absence of Wigglesworthia , tsetse females are sterile, and adult progeny are immune compromised. The functional contributions for Sodalis are less known, while Wolbachia cause reproductive manupulations known as Cytoplasmic Incompatibility (CI). High GpSGHV virus titers result in reduced fecundity and lifespan, and have compromised efforts to colonize flies in the insectary for large rearing purposes. We investigated the within community effects on the density regulation of the individual microbiome partners in tsetse lines with different symbiotic compositions. We show that absence of Wigglesworthia results in loss of Sodalis in subsequent generations possibly due to nutritional dependancies between the symbiotic partners. While an initial decrease in Wolbachia and GpSGHV levels are also noted in the absence of Wigglesworthia , these infections eventually reach homeostatic levels indicating adaptations to the new host immune environment or nutritional ecology. Absence of all bacterial symbionts also results in an initial reduction of viral titers, which recover in the second generation. Our findings suggest that in addition to the host immune system, interdependencies between symbiotic partners result in a highly tuned density regulation for tsetse’s microbiome. Page 225

Publications resulting from this CRP ALAM , U., J. M EDLOCK , C. B RELSFOARD , R. P AIS , C. L OHS et al. , 2011 Wolbachia Symbiont Infections Induce Strong Cytoplasmic Incompatibility in the Tsetse Fly Glossina morsitans. PLoS Pathog 7: e1002415. ATTARDO , G. M., C. L OHS , A. H EDDI , U. H. A LAM , S. Y ILDIRIM et al. , 2008 Analysis of milk gland structure and function in Glossina morsitans: milk protein production, symbiont populations and fecundity. J Insect Physiol 54: 1236-1242. BONOMI , A., F. B ASSETTI , P. G ABRIELI , J. B EADELL , M. F ALCHETTO et al. , 2011 Polyandry is a common event in wild populations of the Tsetse fly Glossina fuscipes fuscipes and may impact population reduction measures. PLoS Negl Trop Dis 5: e1190. DOUDOUMIS , V., G. T SIAMIS , F. W AMWIRI , C. B RELSFOARD , U. A LAM et al. , 2011 Detection and characterization of Wolbachia infections in laboratory and natural populations of different species of tsetse flies (genus Glossina ). BMC Microbiology in press . KARIITHI , H. M., I. A. I NCE , S. B OEREN , A. M. A BD -ALLA , A. G. P ARKER et al. , 2011 The Salivary Secretome of the Tsetse Fly Glossina pallidipes (Diptera: Glossinidae) Infected by Salivary Gland Hypertrophy Virus. PLoS Negl Trop Dis 5: e1371. PAIS , R., C. L OHS , Y. W U, J. W ANG and S. A KSOY , 2008 The obligate mutualist Wigglesworthia glossinidia influences reproduction, digestion, and immunity processes of its host, the tsetse fly. Appl Environ Microbiol 74: 5965-5974. Wang, J., Brelsfoard, C. Wu, Y. and Serap Aksoy , Intercommunity effects on microbiome and GpSGHV density regulation in tsetse flies, JIP Tsetse Symposium, under review WEISS , B., and S. A KSOY , 2011 Microbiome influences on insect host vector competence. Trends Parasitol. WEISS , B., M. M ALTZ and S. A KSOY , 2012 Obligate symbionts activate immune system development in the tsetse fly. Journal of Immunology. WEISS , B. L., J. W ANG and S. A KSOY , 2011 Tsetse immune system maturation requires the presence of obligate symbionts in larvae. PLoS biology 9: e1000619.

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Page 227

Evaluating Wolbachia prevalence, strain-fingerprinting tools, and experimental hybrid

formation in Glossina

Daniela I. Schneider and Wolfgang J. Miller

Laboratories of Genomes Dynamics, Department for Cell and Developmental Biology, Centre of Anatomy and Cell Biology, Medical University of Vienna, Austria

1. Abstract Wolbachia strain typing is routinely based on sequence data derived from the highly dynamic Wolbachia Surface Protein gene ( wsp ), which is prone to strong adaptive evolution and a hotspot for inter-strain recombination. This single locus, however, is not informative in respect to distinguishing CI-inducing from neutral or even mutualistic strain phenotypes. Hence, Wolbachia -specific Multi Locus Strain Typing (MLSTs) and Ankyrin gene (ANKs) marker systems were successfully applied for high resolution phylogenetic strain typing. Novel Wolbachia marker sets cover hypervariable intergenic regions, i.e., variable-number-of- tandem-repeats (VNTRs) and multi-copy insertion sequence (IS) elements. These markers have the potential to be used as an expanded PCR-based multi-locus typing approach, known as Multiple Locus VNTR Analysis (MLVA) that may become an important tool for providing detailed analysis of microbial epidemiology and population genetics. Here, we report on the high applicability of such a novel VNTR-based (Variable-Number-Tandem-Repeat) molecular screening tool for fingerprinting Wolbachia -infections in tsetse flies in Glossina morsitans morsitans, Glossina morsitans centralis , and Glossina brevipalpis. Moreover, we show that certain Wolbachia-infections in Glossina spp. are capable of escaping standard PCR screening methods by ‘hiding’ as low-titer infections below detection threshold. Application of a highly sensitive PCR-blot enhanced symbiont detection limit massively, and allowed us to consequently trace, unequivocally, Wolbachia -infections at high prevalence in laboratory- reared G. swynnertoni individuals. Finally, we highlight the tremendous power of Wolbachia to switch in Glossina inter-species hybrid backgrounds from their natural low or moderate titer levels into levels above these native titers.

2. Background Strain typing of closely related Wolbachia symbionts in insects is challenging and dependent on the choice and information value of marker genes under consideration (recently reviewed in Schneider et al. 2011). Historically, the main body of Wolbachia strain-typing approaches and phylogenies were elaborated on the basis of sequence data derived from the 644-bp sequence of the highly dynamic Wolbachia Surface Protein wsp gene (ZHOU ET AL ., 1998)

Page 228 which is under strong adaptive evolution and a hotspot for interstrain recombination (see

WERREN & BARTOS 2001). In contrast to Wolbachia Ankyrin marker genes (ANKs), the wsp gene is not informative in respect to distinguish CI-inducing from neutral or even mutualistic strain phenotypes (ITURBE -ORMAETXE ET AL ., 2005). Recently, Wolbachia -specific Multi Locus Strain Typing marker systems (MLSTs) were successfully used for phylogenetic strain typing

(BALDO ET AL . 2006; PARASKEVOPOULOS ET AL . 2006). For higher resolution of strain phylogenies and to distinguish even very closely related Wolbachia strains in different host systems we have developed a new set of hypervariable marker systems covering mobile insertion sequences (ISs) and Variable-Number-of-Tandem-Repeats (VNTRs) loci (RIEGLER ET

AL ., 2005; MILLER & RIEGLER 2006; RIEGLER et al. 2012). Hence an expanding arsenal of novel diagnostic Wolbachia marker systems is available now, that have been applied on strain typing of tsetse fly Wolbachia from natural populations and lab colonies.

3. Experimental strategies and major results

3.1 Applying Wolbachia multi-copy IS primer sets for infection diagnostics Mobile genetic elements present universal components of all living organisms and in prokaryotes, they are involved in genomic plasticity, rearrangements and virulence acquisition, and hence, are important elements in bacterial genome evolution. Insertion sequences (ISs) make up the main fraction of prokaryotic transposable elements encoding a transposase gene mediating their transposition , i.e., their ability to move to another locus in a genome and to increase their own copy numbers (CHANDLER & MAHILLON 2002). The complete genome of the sequenced Wolbachia strain wMel of D. melanogaster (WU ET AL ., 2004) occupies at least 50 copies of bacterial ISs and the wSim genome of D. simulans even more that 80 transposable elements (SALZBERG ET AL ., 2005). These numbers are highly unusual for a bacterium and it has been assumed that the mobility of these genomic parasites causing genome dynamics might favor adaptability to their host (WU ET AL ., 2004). All thirteen IS5 transposon copies of wMel have identical sequences suggesting ongoing mobility within this Wolbachia genome. We have recently developed an adaptor-mediated PCR strategy in order to assay IS5 insertion site polymorphism and mobility in wMel for monitoring Wolbachia spreading dynamics and strain replacement in worldwide populations of D. melanogaster over the last 80 years

(RIEGLER ET AL ., 2005). Furthermore, recent mobility of IS5-like elements was detected in a broader range of Wolbachia strains belonging to A- and B-supergroup members (ITURBE -

ORMAETXE ET AL ., 2005; DURON ET AL . 2005; CORDAUX ET AL . 2008). In the course of this CRP we have extracted and designed consensus sequences of five Wolbachia -specific candidate transposons belonging to different IS families by using GenBank

Page 229 database searches and the software program IS-Finder (http://www-is.biotoul.fr/). Based on the hitherto limited number of available Wolbachia genomes at NCBI, for each IS family member we have roughly estimated in silico the individual copy numbers from three complete and four unfinished Wolbachia genomes presently available at NCBI ( Table 1 ). For IS3, IS4 and IS5 elements, significant BLAST similarity hits were only obtained from the four Wolbachia genomes belonging to A-supergroup members whereas no significant hits were detected in the ones from B- and D-supergroup representatives. On the other hand, A-supergroup members presumably lack IS256 elements but in the two B-supergroup representatives wPip PEL and wPip JHB of Culex quinquefasciatus IS256 elements have reached considerable high copy numbers suggesting massive transpositional activities. Finally, the ISNew element, originally isolated by WU ET AL . (2004) from the wMel genome, seems omnipresent in all three super-group representatives with copy numbers ranging between 2 and 10 ( Table 1 ).

Completed genomes Unfinished genomes

wMel (A) wPip (B) wBru (D) wPip (B) wWil (A) wAna (A) wSim (A)

Drosophila melanogaster Culex quinquefasciatus Pel Brugia malayi TRS Culex quinquefasciatus JHB Drosophila willistoni Drosophila ananassae Drosophila simulans IS5 13 0 0 0 2/1 13/6 10/7 IS4 3 0 0 0 2/1 3 3/1 IS3 12/3 0 0 0 7/2 6/8 6/7 IS256 0/2 19/1 0 41/1 0/1 0 0 ISNew 12 2 2/1 2/1 6/2 10/1 5/1

Table 1. Approximation of individual IS copy numbers determined by in silico analysis of presently available Wolbachia genomes at NCBI. Numbers give estimates of full-sized IS copies per genome, whereas the ones post-slash indicate copy numbers of truncated IS elements.

Based on wsp sequence data obtained from literature, all Wolbachia strains so far isolated from tsetse fly samples belong to A-supergroup (CHENG ET AL . 2000). Hence we assume that at least four of our five IS elements might be appropriate multi-copy candidates for further IS- diagnostics and fingerprinting of Wolbachia in Glossinas spp. Wolbachia -specific consensus primer sets were designed particularly targeting IS3, IS4, IS5, IS256 and ISnew elements. The design of the five IS-specific consensus primer sets has been performed based on BLAST

Page 230 similarity searches, multiple sequence alignments and sectional comparisons. Using these new primer sets, we have screened DNA derived from lab colonies of G. morsitans centralis, G. brevipalpis and G. palpalis, kindly provided from Seibersdorf (Figure 1 ). Further improvement of the primer design of individual IS elements will be feasible as soon as more partial or even complete genome sequences from different Wolbachia strains will be accessible in the near future. The availability of the first complete tsetse Wolbachia genome from G. morsitans morsitans will further improve the design and quality of these IS primer sets.

Figure 1. IS-element PCR on Glossina species. Gbr Gmc Gmc Gpal Gpp Gbr Gmc M Gbr Gmc M Gbr Gmc DNA samples deriving from G. brevipalpis (Gbr ), G. m. centralis (Gmc ), G. pallidipes (Gpal ), and 1000 750 G. palpalis (Gpp ) were tested for Wolbachia 500 infection. M = DNA ladder. IS3 IS4 IS5 IS256 ISnew

In our IS-element screen, IS3, IS4 as well as ISnew turned out the most sensitive marker for detecting Wolbachia in different Glossina species, whereas IS5 and IS256 do not work with the sensitivity. IS4 seems suited very well to detect rather low titer Wolbachia like determined in G. brevipalpis .

3.2 Tracking Wolbachia low-titer infections with a highly sensitive PCR-blot technique Application of the IS multi-copy PCR approach is of great value for assessing Wolbachia infections in Glossina by enhancing detection sensitivity. Some natural Wolbachia infections within this genus, however, persist at densities far below PCR detection limit and are thus overlooked even with the multi-copy PCR approach. We have hence applied a more sensitive detection tool based on a combination of standard PCR and hybridization, and tracked extremely low titer Wolbachia infections in G. swynnertoni . No Wolbachia was detected in samples from this subspecies via standard wsp -PCR ( Figure 2A ). By applying our PCR-blot technique to the same sample set however, we significantly enhanced the detection capacity of our method and more samples tested positive for Wolbachia -infection. In G. swynnertoni 3/3 instead of 0/3 individuals and in 4/4 G. m. morsitans individuals were positive instead of only 2/4 ( Figure 2B ). Via employing our highly sensitive detection tools, i.e., PCR-blot technique and VNTR-based infection screen, to more samples from each tsetse species (data not shown) we estimated Wolbachia-prevalence as follows: G. m. centralis 57/58 (98%), G. m. morsitans 33/40 (83%); G. swynnertoni 7/10 (70%), and G. brevipalpis 9/13 (70%) respectively. These data suggest a generally high fixation rate of 70% on average in all tested Glossina species.

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Figure 2 . Detection of low-titer Wolbachia Gsw Gmc Gmm Gbr - in tsetse flies (A) Standard wsp -PCR on Glossina spp. (B) same gel after PCR-blot probed with a digoxygenin-labeled wsp - fragment (Arthofer et al. 2009). According to wsp -PCR, Wolbachia -infections are high in G. m. centralis (Gmc ), intermediate in G. m. morsitans (Gmm ), low in G. brevipalpis (Gbr ), but the PCR-blot technique significantly enhances detection sensitivity, especially in G. m. morsitans , G. brevipalpis and G. swynnertoni . Neg. control = Wolbachia - uninfected D. simulans, STC .

3.3 Isolating novel hyper-variable marker systems for tsetse fly Wolbachia fingerprinting by applying the Variable Number of Tandem Repeats (VNTRs) marker system.

Repetitive DNA, which occurs in large quantities in eukaryotic cells, has been increasingly identified in prokaryotes (reviewed in VAN BELKUM ET AL . 1998). Microbial genomes contain a variety of repetitive DNA sequences, accounting for up to 5% of the genome (USSERY ET AL ., 2004). Most of these repetitive DNA elements are of unknown function and are infrequently associated with coding regions and consequently are located in both intergenic and extragenic regions of the microbial genome. Multiple-locus variable-number-of-tandem-repeat (VNTR) analysis is a highly successful method for studying genetic variability of many bacterial species, especially pathogens (KEIM ET AL . 2000; KLEVYTSKA ET AL . 2001; LIAO ET AL . 2006; and reviewed in TOP ET AL . 2004; LINDSTEDT 2005). Indeed, VNTRs have been proven to provide a high level of discriminatory power for strain differentiation because of their high mutability prone by replication slippage and ectopic recombination between cluster units ( VAN BELKUM ET

AL . 1998).

On the base of the complete genome sequence of Wolbachia from D. melanogaster wMel (WU

ET AL ., 2004), a novel set of hyper-polymorphic markers became available (RIEGLER ET AL ., 2005). The isolation and successful application of Wolbachia -specific satellite loci composed of VNTRs allowed us to discriminate between at least five distinctive Wolbachia strain variants within D. melanogaster (RIEGLER ET AL ., 2005) thought to be a single infection based on wsp sequence. This outlines the limitations of the wsp marker system for Wolbachia strain typing and the urgent need for better and more sensitive molecular marker systems in order to separate closely related Wolbachia strains causing distinctive phenotypes. By using the program Tandem Repeats Finder 3.21 (BENSON , 1999) we have detected 63 sites with direct tandem repeats in the wMel chromosome, having period sizes from 10bp to 291bp and

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internal match from 72 to 100% (RIEGLER ET AL ., 2005; MILLER & RIEGLER 2006). PCR primer sets were designed to the flanking regions of the repeats and applied on different samples of insect species infected with closely related or identical Wolbachia strains according to their wsp sequence. In the case of the primer set VNTR-141 amplicon size differences were found ranging from 0.3 to 1.2-kb harboring from zero ( wWil of D. willistoni ) to seven copies ( wMel of D. melanogaster ) of the 141-bp repeat. For example, Wolbachia infections of D. willistoni and D. simulans share 100% identity at wsp and various ANK loci but differ in tissue tropism in their respective hosts (MILLER & RIEGLER 2006). Whereas the VNTR-141 locus of wAu in D. simulans harbors 1.7 units of the 141-bp core repeat, wWil shows an ancestral state of the locus composed of a limited number of short sub-repeats only ( Figure 3 ).

Figure 3. Organization of the VNTR-141 locus in Glossina spp. The 464-bp fragments of wGmm , wGmc , wGsw , and wGbr share a small 8-bp insertion between blue and yellow repeat box and a 104-bp insertion (grid) at the 3’-end of the locus; the 347-bp chromosomal fragment of Gmm and the 483-bp wGbr fragment lack both. Green and blue boxes represent 15-bp repeats, yellow and red ones 13-bp repeats. The Gray arrow represents the complete core 108-bp repeat of wAu from D. simulans . Abbreviations: wGmm, wGmc, wGbr, and wGsw symbolize Wolbachia from G . m. morsitans , G. m. centralis , G. brevipalpis , and G. swynnertoni , respectively. Black arrow indicates the 141-bp master unit from wMel-like strains.

To assess full Wolbachia -diversity in tsetse flies from laboratory strains and field collections, we have applied a robust and highly sensitive VNTR-based screen on three members of the morsitans group, i.e ., G. m. morsitans , G. m. centralis and G. swynnertoni , as well as on G. brevipalpis belonging to the distantly related fusca group ( Figure 4 ). VNTR-141 PCR on laboratory and field samples of G. m. morsitans revealed two fragments of diagnostic length (347-bp and 464-bp, respectively; see Figure 4A , lanes a), whereas Wolbachia from G. m.

Page 233 centralis displayed only one amplicon of 464-bp (lane b). In Wolbachia from G. swynnertoni , the VNTR primer set amplifies a fragment similar in size to G. m. centralis Wolbachia (464-bp), but of weaker intensity (lane c). Finally the G. brevipalpis infection is characterized by a VNTR- 141 fragment of unique size of 483-bp (lane d).

Figure 4 . (A) VNTR-141-PCR on four Glossina species. Samples are either laboratory-reared or field collected single individuals of (a) G . m. morsitans Gmm , (b) G. m. centralis Gmc , (c) G. swynnertoni Gsw , and (d) G. brevipalpis Gbr. Black arrowheads indicate a 347-bp fragment in Gmm of chromosomal origin; a 464-bp fragment of cytoplasmic origin in Gmm , Gmc , and Gsw ; and a larger 483-bp fragment in Gbr . (B) VNTR-141- PCR on wild type (wt) and apo-symbiotic Gmm. The cytoplasmic 464-bp band is more prominent in wt, whereas the nuclear 347-bp band is strong in the aposymbiotic Gmm . DNA quality was assessed via 12S rRNA-specific PCR.

The finding of two diagnostic VNTR-141 bands in all G. m. morsitans samples suggests either the existence of a double infection of the endosymbiont in G. m. morsitans , a duplication of the VNTR locus on the G. m. morsitans Wolbachia chromosome, or the horizontal transfer of the locus onto the host chromosome. Nuclear translocation events of Wolbachia genes have been reported in a variety of insect and nematode hosts ( REVIEWED IN BLAXTER 2007). Indeed,

DOUDOUMIS ET AL . (2012) reported recently on nuclear copies of at least three Wolbachia genes (16S rRNA, wsp and fbpA ) in G. m. morsitans chromosomes. We hence determined the origin of the two VNTR-141 fragments via diagnostic PCR on wild type (wt) and antibiotic- treated (apo-symbiotic; DNA kindly provided by S. Aksoy) G. m. morsitans samples. As shown in Figure 4B, intensity of the upper VNTR band in wt flies is significantly decreased upon antibiotic treatment, suggesting a cytoplasmic origin, and the persistence of the smaller fragment in apo-symbiotic samples points towards a nuclear localization.

3.4 Wolbachia titer dynamics in Glossina inter-species hybrids Recent, independent studies reported on the enormously high importance of symbiont replication control, host background, and maintenance of a certain balance between host and symbiont loads (POINSOT ET AL ., 1998; MCGRAW ET AL . 2002; VENETI ET AL . 2004; BORDENSTEIN

ET AL . 2006; MILLER ET AL . 2010; CHAFEE ET AL . 2011; LOGIN ET AL . 2011; SERBUS ET AL . 2008 AND 2011). In the Drosophila paulistorum species complex, backgrounds from inter-semispecies hybrids negatively influence host fitness and fecundity massively by boosting Wolbachia -titer

(MILLER ET AL . 2010). Furthermore a two-fold titer increase of native Wolbachia was reported

Page 234 recently in the F1 hybrids between closely related parasitoid wasps of the genus Nasonia

(CHAFEE ET AL . 2011). Here we generated inter-species hybrids between members of the Glossina morsitans group and assessed their Wolbachia -infection titer in comparison to the corresponding parental generation. By taking advantage of our PCR-blot technique, we revealed a dramatic increase of Wolbachia -titer in all hybrid backgrounds of Gmm x Gmc (the reciprocal cross does not produce living offspring); Gsw x Gmc , and Gmc x Gsw ; Gmm x Gsw , and Gsw x Gmm , compared to their corresponding parents (data not shown). In order to determine Wolbachia -titer levels in parents and hybrids in a quantitative way, we performed Wolbachia - specific 16S rRNA qRT-PCR. Three-day old female and male F1 hybrids were analyzed for Wolbachia -infection level and compared to symbiont load in the corresponding three-day old individuals from the parent colonies. Compared to PCR-hybridization analyses, qRT-PCR data support our finding of massive Wolbachia -titer increase in all the five hybrid backgrounds.

In summary we have succeeded to establish novel and highly sensitive molecular tools and protocols for diagnosing Wolbachia prevalence, strain diversity, plus symbiont titer dynamics in native tsetse flies as well as in their respective inter-species hybrids, and finally, to decipher the existence of “hidden” low titer infections in samples that were considered as uninfected by means of standard PCR approaches. A research paper has been recently submitted to the Journal of Invertebrate Pathology which will present our innovative detection tools and novel findings on Wolbachia -tsetse fly symbiosis that will be - as we think – highly informative for our colleagues and will help to understand the entire biological complexity in this multi-factor symbiosis.

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