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Focus 19 Trees, A.J. et al. (1993) Prevalence of antibodies to Neospora caninum in a population of urban dogs in England. Vet. Rec. 132, 125–126 20 Schares, G. et al. (1998) The efficiency of vertical transmission of Neospora caninum in dairy cattle analysed by serological techniques. Vet. Parasitol. 80, 87–98 21 Osawa, T. et al. (1998) A multiple antigen ELISA to detect Neospora specific antibodies in bovine sera, bovine foetal fluids, ovine and caprine sera. Vet. Parasitol. 79, 19–34 22 Björkmann, C. et al. (1994) Neospora caninum in dogs: detection of antibodies by ELISA using an iscom antigen. Parasite Immunol. 16, 643–648 23 Björkmann, C. et al. (1997) An indirect enzyme linked immunoassay (ELISA) for demonstration of antibodies of Neospora caninum in serum and milk of cattle. Vet. Parasitol. 68, 251–260 24 Williams, D.J.L. et al. (1997) Novel ELISA for detection of Neospora-specific antibodies in cattle. Vet. Rec. 140, 328–331 25 Packham, A.E. et al. (1998) A modified agglutination test for Neospora caninum – development, optimisation, and comparison to the indirect fluorescent-antibody test and enzyme linked immunosorbent assay. Clin. Diagn. Lab. Immunol. 5, 467–473 26 Williams, D.J.L. et al. Evaluation of a commercial ELISA to detect serum antibody to Neospora caninum in cattle. Vet. Rec. (in press) 27 Paré, J. et al. (1995) Interpretation of an indirect fluorescent antibody test for diagnosis of Neospora sp. infection in cattle. J. Vet. Diagn. Invest. 7, 273–275 28 Björkman, C. and Uggla, A. (1999) Serological diagnosis of Neospora caninum infection. Int. J. Parasitol. 29, 1497–1507 29 Sasai, K. et al. (1998) A chicken anti-conoid monoclonal antibody identifies a common epitope which is present on motile stages of Eimeria, Neospora and Toxoplasma. J. Parasitol. 84, 654–656 30 Schares, G. et al. (1999) Serological differences in Neospora caninum-associated epidemic and endemic abortions. J. Parasitol. 85, 688–694

31 Wouda, W. et al. (1998) Serodiagnosis of neosporosis in individual cows and dairy herds, a comparative study of three enzyme-linked immunosorbent assays. Clin. Diagn. Lab. Immunol. 5, 711–716 32 Paré, J. et al. (1997) Neospora caninum antibodies in cows during pregnancy as a predictor of congenital infection and abortion. J. Parasitol. 83, 82–87 33 Anderson, M.L. et al. (1997) Evidence of vertical transmission of Neospora sp. infection in dairy cattle. J. Am. Vet. Med. Assoc. 210, 1169–1172 34 Davison, H.C. et al. (1999) Herd-specific and age specific seroprevalence of Neospora caninum in 14 British dairy herds. Vet. Rec. 144, 547–550 35 Barber, J.S. and Trees, A.J. (1996) Clinical aspects of 27 cases of neosporosis in dogs. Vet. Rec. 139, 439–443 36 Dubey, J.P. et al. (1997) Antibody responses of cows during an outbreak of neosporosis evaluated by indirect fluorescent antibody test and different enzyme-linked immunosorbent assays. J. Parasitol. 83, 1063–1069 37 Schares, G. et al. (1999) Bovine neosporosis: comparison of serological methods using outbreak sera from a dairy herd in New Zealand. Int. J. Parasitol. 29, 1659–1667 38 Thurmond, M. and Hietala, S. (1995) Strategies to control Neospora infection in cattle. Bovine Practitioner 29, 60–63 39 Cox, B.T. et al. (1998) Serology of a Neospora abortion outbreak on a dairy farm in New Zealand: a case study. N. Z. Vet. J. 46, 28–31 40 Barber, J.S. et al. (1997) Prevalence of antibodies to Neospora caninum in different canid populations. J. Parasitol. 83, 1056–1058 41 Paré, J. et al. (1996) Congenital Neospora caninum infection in dairy cattle and associated calfhood mortality. Can. J. Vet. Res. 60, 133–139 42 Björkmann, C. et al. (1996) Neospora species infection in a herd of dairy cattle. J. Am. Vet. Med. Assoc. 208, 1441–1444 43 Harper, P.A.W. (1999) Are your cattle aborting? Agfact AO. 9. 58, 1st edn 1994. New South Wales Agriculture 44 McAllister, M.M. (1999) Uncovering the biology and epidemiology of Neospora caninum. Parasitol. Today 15, 216–217

Tsetse – A Haven for Microorganisms S. Aksoy Arthropods are involved in the transmission of parasitic and viral agents that cause devastating diseases in animals and plants. Effective control strategies for many of these diseases still rely on the elimination or reduction of vector insect populations. In addition to these pathogenic organisms, arthropods are rich in microbes that are symbiotic in their associations and are often necessary for the fecundity and viability of their hosts. Because the viability of the host often depends on these obligate symbionts, and because these organisms often live in close proximity to disease-causing pathogens, they have been of interest to applied biologists as a potential means to genetically manipulate populations of pest species. As knowledge on these symbiotic associations accumulates from distantly related insect taxa, conserved mechanisms for their transmission and evolutionary histories are beginning to emerge. Here, Serap Aksoy summarizes current knowledge on the functional and evolutionary biology of the multiple symbionts harbored in the medically and agriculturally important insect group, tsetse, and their potential role in the control of trypanosomiasis. Tsetse (Diptera: Glossinidae) are the vectors of African trypanosomes, the causative agents of sleeping sickness Serap Aksoy is at the Department of Epidemiology and Public Health, Section of Vector Biology, Yale University School of Medicine, 60 College St., 606 LEPH, New Haven, CT 06510, USA. Tel: +1 203 737 2180, Fax: +1 203 785 4782, e-mail: [email protected] 114

disease in humans, as well as various diseases in animals. Recently, tsetse-transmitted diseases have been on the rise, causing severe economic hardships in many already stressed communities. The extensive antigenic variation the parasites display in their mammalian host has hampered efforts to develop effective vaccines, and disease management strategies currently rely on the treatment of infected hosts by chemotherapy and on reduction of tsetse challenge by eradication or suppression approaches. However, the success of active surveillance attempts requires the development of improved methods with greater diagnostic sensitivities than the traditional detection techniques1. The trypanocidal drugs used for treatment are expensive and highly toxic, with adverse side effects; in addition, their efficacy has recently been challenged in the presence of increasing parasite drug resistance detected in patients2. Efforts to control vector populations have been based largely on ground and aerial spraying, as well as the direct application of insecticides to cattle in farming communities. Recently, traps and targets have been used extensively to reduce vector challenge, but reports on their sustainability have been mixed because their long-term success relies heavily on community participation, awareness and resources. In geographically isolated areas, such as the island of Zanzibar, eradication attempts using the ‘sterile insect technique’ (SIT) have been successful, and have resulted

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Focus in complete elimination of disease from farming communities. On the mainland, however, this approach requires area-wide participation to be effective, as well as geographically isolated populations to prevent reinvasion of cleared lands. Because each individual methodology has its limitations, there is growing interest in integrated control programs that can incorporate several different schemes, depending on the ecology and epidemiology of the disease, and on the cultural preferences and resources of each community3. Given the recent advances in molecular technologies, there has been an effort to develop alternative genetic-based strategies that can be included in the battery of tools available for integrated control programs. As one such approach, transgenesis is being explored – it aims to modulate the vector competence of insects; ie. to eliminate the ability of insects to transmit disease agents by introducing and expressing foreign genes with antipathogenic properties to interfere with pathogen viability, development and/or transmission. These pathogen-refractory insects can then replace their natural susceptible counterparts by virtue of mechanisms that would confer reproductive advantages to the engineered insects4,5. Towards this end, it has been possible to introduce foreign genes into several different insects by traditional transformation vectors (transposable elements or viral-transducing agents)6. In addition to this direct germ-line transformation approach, foreign genes have also been expressed in the symbiotic microorganisms that are closely associated with several arthropods, including tsetse4. Characterization of tsetse symbionts Many insects that have limited diets (such as blood, plant sap or wood) throughout their entire developmental cycle rely on microbial symbionts for additional nutrients not found in their restricted diet, and which they are unable to synthesize7,8. Microorganisms with different ultrastructural characteristics have also been reported from various tissues of tsetse, including midgut, hemolymph, fat body and ovaries9–13. Until recently, their taxonomic identification was unknown; however, PCR-based phylogenetic studies have now shown that they represent three distinct organisms14. Two of these organisms are present in gut tissue: the primary (P)-symbiont (Wigglesworthia glossinidia) resides intracellularly in the specialized epithelial cells (bacteriocytes), which form a U-shaped organ (bacteriome) in the anterior gut15,16, while the secondary (S)symbiont (Sodalis glossinidius) is present in midgut cells17–19. The third organism, which has been characterized from reproductive tissue, is related to Wolbachia pipientis14. Tsetse are viviparous, retaining each egg within a uterus, where it hatches after fertilization; one young larva matures and is finally expelled as a fully developed third instar larva. Each female mates once and can deposit 5–7 offspring during its 3–4 month lifespan. During its intrauterine life, the larva receives nutrients and both of the gut symbionts from its mother via milk-gland secretions20,21. The Wolbachia bacterium is present in the ovaries and is transovarially transmitted through maternal lineages. Given the unique reproductive biology of tsetse, however, all three symbionts are, in essence, maternally acquired by the progeny. It is difficult to study the individual functions of the multiple symbionts in tsetse. Attempts to eliminate the Parasitology Today, vol. 16, no. 3, 2000

symbionts by administration of antibiotics, lysozyme and specific antibodies result in retarded growth of the insect and a decrease in egg production, preventing the ability of the aposymbiotic host to reproduce22–24. The ability to reproduce, however, can be partially restored when the aposymbiotic tsetse receive a bloodmeal that is supplemented with B-complex vitamins (thiamine, pantothenic acid, pyridoxine, folic acid and biotin), suggesting that the endosymbionts play a role in metabolism that involves these compounds25. In addition to their role in nutrition, there has been indirect evidence suggesting that the presence of symbiont(s) might also enhance the establishment of trypanosome infections in the midgut26. It has been shown that Sodalis produces at least one type of chitinase enzyme, which might be responsible for increasing the trypanosome susceptibility of its host insect27,28. It has been possible to culture Sodalis in vitro29,30, and the availability of this culture system has permitted the biochemical characterization of this organism. Carbon substrate assimilation tests suggest that Sodalis primarily utilizes N-acetylglucosamine and raffinose as its primary carbon sources in vitro19. The use of N-acetylglucosamine might reflect an adaptation by the organism to the gut environment of its host, which is largely made of chitin (polymerized N-acetyl-D-glucosamine). The W. pipientis-like symbiont detected in tsetse gonads has been found to infect a wide range of invertebrate hosts. In one survey in the neotropics, more than 15% of the analyzed taxa were reported to carry this group of microorganisms31. In the hosts they infect, Wolbachia have been shown to cause a variety of reproductive abnormalities, one of which is termed cytoplasmic incompatibility (CI), and when expressed, often results in embryonic death owing to disruptions in early fertilization events32. In an incompatible cross, the sperm enters the egg but does not successfully contribute its genetic material to the potential zygote. In most species, this results in very few hatching eggs. The infected females have a reproductive advantage over their uninfected counterparts as they can produce successful progeny with both the imprinted and normal sperm. This reproductive advantage allows the infected insects to spread into populations. For functional studies, it has been possible to cure most insects of their Wolbachia infections by administering antibiotics in their diet; however, this approach has not been feasible in tsetse because the antibiotic treatment of flies results in the clearing of all bacterial symbionts, including the nutritional obligatory associations described above, and in the absence of these gut symbionts the flies become sterile. The analysis of tsetse laboratory colonies has shown that 100% of sampled individuals carry Wolbachia infections, making it impossible to investigate Wolbachia-mediated effects by traditional mating experiments using infected and uninfected individuals. The prevalence of infections in field populations, however, has shown that various tsetse species have been infected with Wolbachia and significant polymorphism exists in the field33. In one survey, although a 98% level of infection was seen in Glossina austeni sampled from South Africa, only 48% of the sampled Kenyan population was found to be infected33. Hence, Wolbachia-infected and -uninfected lines can be developed from these heterogeneous field populations to elucidate the functional role of this 115

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G. austeni -S Fig. 1. The phylogenetic placement of Glossina species and their primary (P)- and secondary (S)-symbionts. The phylogenetic tree constructed by maximum parsimony based on the internal transcribed spacer 2 (ITSG. fuscipes -S 2) sequence of the indicated Glossina species (a). The analysis included 408 sites (432 with gaps), of which 222 (246 with gaps) were variable and 88 (112 with gaps) were informative. One representative tree is shown (tree length 5 292; CI 5 0.897). The bootstrap confidence values (300 replications) are presented G. brevipalpis -S at the nodes, the values in parentheses are bootstrap values where gaps have been included as weighted characters; numbers below denote branch length values. Drosophila yakuba and Musca domestica sequences S. zeamais -S are used as outgroups. The grouping of the palpalis, morsitans, austeni and fusca species is indicated. F denotes analysis done using field-collected material; C denotes colony material; numbers refer to independent PCR amplification, cloning and sequencing experiments. Phylogenetic analysis of Glossina symbionts based on their 16S rDNA sequence analysis (b). P denotes primary symbiont Wigglesworthia, and S denotes the secondary symbionts analyzed from the corresponding species of Glossina. Escherichia coli was used as the outgroup. The analysis included 893 sites of which 205 were variable and 121 were informative. One representative tree is shown (tree length 5 216; CI 5 0.843; bootstrap values for 500 replications are shown at the nodes). Abbreviations: CI, Consistency index; G. m. morsitans, Glossina morsitans morsitans; Glossina p. palpalis, G. palpalis palpalis. (Adapted with permission, from Ref. 35.)

organism in tsetse biology. A phylogenetic analysis of the Wolbachia strain types infecting different species of tsetse has shown that they are different and, as such, represent independent acquisitions33. Evolutionary histories of symbionts Because it has been difficult to cultivate many of these fastidious, and often intracellular organisms in vitro, their correct taxonomic positioning has been controversial. Recent advances in PCR-based technologies, as well as the use of nucleic acid sequences in phylogenetic reconstructions, have now provided additional insight into the relationships among bacteria34. The 16S rDNA sequence-based characterization of tsetse gut organisms indicate that they are members of the family Enterobacteriaceae. The analysis of Wigglesworthia and Sodalis from species representing the four subgenera of Glossina: fusca, morsitans, palpalis and austeni, has shown that they each form a distinct lineage in the g-subdivision of the Proteobacteria16 (Fig. 1). The phylogenetic relationship of the different Glossina host species has also been determined independently using the DNA sequence of the internal transcribed spacer 2 116

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(ITS-2) regions of the rDNA locus35. The 16S rDNA sequence of Wigglesworthia from different species of tsetse and the ITS-2 sequence of tsetse species have been found to be variable (5–10%) and, hence, were used for phylogenetic reconstruction of the two histories. When tsetse and Wigglesworthia phylogenies were compared, they were found to display identical relationships among the different species. This finding suggests that a tsetse ancestor was infected with a bacterium, and from this ancestral pair evolved the tsetse host and endosymbiont, comprising the species of tsetse and Wigglesworthia strains that exist today and overruling possible horizontal transfer events between species. The bacteriome-associated P-symbionts of other insect taxa such as aphids36, whiteflies37, mealybugs38, cockroaches39 and carpenter ants40 have similarly been shown to represent distinct lineages that parallel the evolutionary histories of their host insect species. Thus, although the microorganisms represent distant taxa in the Eubacteria, their evolutionary associations with their insect hosts are strikingly similar. The genome size of Buchnera, the P-symbiont of aphids, is 650-kb41, and the size of Wigglesworthia is also in this Parasitology Today, vol. 16, no. 3, 2000

Focus range (S. Aksoy, unpublished), both much smaller than their closely related, free-living relatives in the Enterobacteriaceae. The small sizes of their genomes might be suggestive of their ancient intracellular histories. Unlike the concordant relationship Wigglesworthia displays with its host, isolates of Sodalis from different tsetse have almost identical 16S rDNA sequences, suggesting that they might either represent recent independent acquisitions by each host species, or alternatively, there may have been multiple horizontal transfer events between species (Fig. 1)21,33. In fact, comparative analysis of the 16S rDNA sequence of Sodalis with other bacteria indicates a close relationship with the symbionts of Sitophilus zeamais and Acyrthosiphon pisum, indicating that this group of microorganisms might share a recent ancestor within the Enterobacteriaceae. Tissue tropism of symbionts A PCR-based assay has been developed with symbiont-specific amplification primers to investigate the tissue tropism of the multiple symbionts in tsetse. Using this assay, Wigglesworthia is found exclusively within the bacteriome-tissue, whereas Sodalis is detected in midgut, muscle, fatbody, hemolymph, milkgland and in salivary glands of certain species18. Furthermore, the density of infections with Sodalis varies in the different species analyzed. Whereas infections in G. morsitans and G. palpalis midgut tissues are maintained at high density, infections in G. austeni and G. brevipalpis are significantly less dense, as measured by both PCR18 and microscopy42. The factors that control tissue tropism and the density of symbionts in insects are not known. Although infections with Wolbachia were initially thought to be largely restricted to the germ-line tissue of their insect hosts, they have recently been shown to be present in a variety of somatic tissues in several different insects43,44. Analysis of the tissue tropism of Wolbachia in tsetse indicate that, whereas in G. morsitans and G. brevipalpis infections are restricted to gonads, in G. austeni they can be detected in various somatic tissues, even in two-day-old teneral flies33. The extensive infections associated with G. austeni somatic tissues might reflect the particular phenotype of the bacterial strain harbored in this species, similar to that reported in the Wolbachia popcorn strain characterized from Drosophila melanogaster44. It remains to be seen whether such an invasive tissue tropism in G. austeni can also result in reduced fitness effects similar to those seen in D. melanogaster44. Symbionts as expression vectors There is no direct germ-line DNA transformation system available for tsetse and the viviparous nature of its reproductive biology makes transgenic approaches that rely on egg microinjections difficult. However, it has been possible to culture the S-symbiont Sodalis in vitro29,30, and a genetic transformation system has been developed so that foreign genes can be introduced and expressed in these cells30. The in vitro-manipulated recombinant Sodalis can be acquired successfully by the intrauterine progeny by means of microinjection into the female parent hemolymph. The recombinant symbionts are passed on to the F1 as well as their offspring, where they synthesize the marker gene product, green fluorescent protein (GFP)18. Because Sodalis is found in close proximity to the site of procyclic trypanosome Parasitology Today, vol. 16, no. 3, 2000

differentiation and replication in the midgut, the synthesis and secretion of antitrypanosomal gene products in these symbionts in vivo can provide a mechanism to disrupt parasite establishment in the gut. However, for the symbiont-based transformation approach to be successful, a significant proportion of the natural symbiont population of the gut will need to be reconstituted by its recombinant counterpart so that foreign gene products can accumulate in the gut to levels where they can interfere with trypanosome biology. The eventual replacement of parasite-susceptible vector populations with engineered refractory flies could provide an additional strategy to reduce disease in the field5. Because Wolbachia infections in tsetse appear to express strong CI phenotypes, the two symbiotic systems could be coupled to drive the phenotypes conferred by the engineered gut symbionts into the field5. This should be feasible because Wolbachia infections rapidly invade populations by virtue of the CI phenomenon they confer, and therefore, they can drive other maternally inherited elements such as mitochondria45 or the engineered gut symbionts into that same population46. Alternatively, the presence of Wolbachia infections in the somatic tissues of various insects, including tsetse, now opens up the possibility of expressing antipathogenic genes directly in this bacterium, which could then replace the susceptible insect populations in the field33. Although no naturally occurring infectious transfer of Wolbachia has been observed, it has become increasingly common to transfer Wolbachia experimentally between different hosts, and even into insects with no prior infection history, making it an attractive gene expression system with a naturally associated driving mechanism47–50. Prospects Using a similar symbiont-based insect transformation approach, it has been possible to block the transmission of Trypanosoma cruzi in Rhodnius prolixus in vivo by expressing an anti-parasite peptide, cecropin A, in its symbiont, Rhodococcus rhodnii in hindgut51. In the R. prolixus system, it has been possible to generate bugs free from their natural gut symbiont flora and to introduce recombinant organisms into these aposymbiotic first-instar animals through artificial coprophagy. In aphids, the facultative S-symbiont (PASS), which is closely related to Sodalis, has been introduced by microinjection from Acyrthosiphon pisum into A. kondoi Shinji (blue alfalfa aphid), as well as into A. pisum-negative clones, where it is maintained in the progeny of the injected mother aphids with a high rate of maternal transmission52. The availability of in vitro symbiont culture and the relative ease of DNA transformation systems in bacteria will facilitate the development of similar disease intervention strategies in other medically and agriculturally important vector systems such as ticks, mites, bed bugs, lice, some species of fleas, planthoppers, white-flies and termites where these associations have been firmly documented. The success of symbiont-based transgenic strategies in insects relies on a good understanding of the molecular and developmental biology of the symbionts, as well as the pathogens transmitted by each system, so that genes with transmission-blocking activities can be identified and expressed efficiently in the correct tissues, thereby adversely affecting pathogen viability. In addition to questions regarding technical success and efficacy, there 117

Focus are many questions about the safety and regulatory concerns for release of genetically modified insects, especially human-biting vectors. Before any release studies can be entertained with recombinant animals, information on environmental and ecological hazards associated with the releases and potential public health risks will need to be deliberated. The successful replacement of vector populations with engineered insects will also depend on the availability of a sound knowledge base of the natural ecology and population biology of these insect species. Acknowledgements Thanks to Scott O’Neill, Leyla Akman, Liangbiao Zheng and the anonymous reviewers for critical reading of the manuscript and thoughtful suggestions. References 1 Rebeski, D. et al. (1999) Improved methods for the diagnosis of African trypanosomosis. Mem. Inst. Oswaldo Cruz 94, 249–253 2 Mulugeta, W. et al. (1997) Long-term occurrence of Trypanosoma congolense resistant to diminazene, isometamidium and homidium in cattle at Ghibe, Ethiopia. Acta Trop. 64, 205–217 3 Politi, C. et al. (1995) Cost-effectiveness analysis of alternative treatments of African gambiense trypanosomiasis in Uganda. Health Econ. 4, 273–287 4 Beard, C. et al. (1998) Bacterial symbiosis in arthropods and the control of disease transmission. Emerg. Infect. Dis. 4, 581–591 5 Beard, C. et al. (1994) Modification of arthropod vector competence via symbiotic bacteria. Parasitol. Today 9, 179 6 Ashburner, M. et al. (1998) Prospects for the genetic transformation of arthropods. Insect Mol. Biol. 7, 201–213 7 Douglas, A.E. (1989) Mycetocyte symbiosis in insects. Biol. Rev. Camb. Philos. Soc. 64, 409–434 8 Buchner, P. (1965) Endosymbiosis of Animals with Plant Micro-organisms, Interscience Publishers 9 Pinnock, D.E. and Hess, R.T. (1974) The occurrence of intracellular rickettsia-like organisms in the tsetse flies, Glossina morsitans, G. fuscipes, G. brevipalpis and G. pallidipes. Acta Trop. 31, 70–79 10 Roubaud, B. (1919) Les particularites de la nutrition et de la vie symbiotique chez les mouches tsetse. Ann. Inst. Pasteur Microbiol. 33, 489–537 11 Shaw, M.K. and Moloo, S.K. (1991) Comparative study on Rickettsia-like organisms in the midgut epithelial cells of different Glossina species. Parasitology 102, 193–199 12 Stuhlman, F. (1907) Beitrage zur kenntnis der Tsetse fliege. Arb. Gesundh. Amte (Berlin) 26, 301–308 13 Wigglesworth, V.B. (1929) Digestion in the tsetse fly: a study of the structure and function. Parasitology 21, 288–321 14 O’Neill, S.L. et al. (1993) Phylogenetically distant symbiotic microorganisms reside in Glossina midgut and ovary tissues. Med. Vet. Entomol. 7, 377–383 15 Aksoy, S. (1995) Wigglesworthia gen. nov. and Wigglesworthia glossinidia sp. nov., taxa consisting of the mycetocyte-associated, primary endosymbionts of tsetse flies. Int. J. Syst. Bacteriol. 45, 848–851 16 Aksoy, S. et al. (1995) Mycetome endosymbionts of tsetse flies constitute a distinct lineage related to Enterobacteriaceae. Insect Mol. Biol. 4, 15–22 17 Aksoy, S. (1995) Molecular analysis of the endosymbionts of tsetse flies: 16S rDNA locus and over-expression of a chaperonin. Insect Mol. Biol. 4, 23–29 18 Cheng, Q. and Aksoy, S. (1999) Tissue tropism, transmission and expression of foreign genes in vivo in midgut symbionts of tsetse flies. Insect Mol. Biol. 8, 125–132 19 Dale, C. and Maudlin, I. (1999) Sodalis gen. nov. and Sodalis glossinidius sp. nov., a microaerophilic secondary endosymbiont of the tsetse fly Glossina morsitans morsitans. Int. J. Syst. Bacteriol. 49, 267–275 20 Ma, W-C. and Denlinger, D.L. (1974) Secretory discharge and microflora of milk gland in tsetse flies. Nature 247, 301–303 21 Aksoy, S. et al. (1997) Phylogeny and potential transmission routes of midgut-associated endosymbionts of tsetse (Diptera: Glossinidae). Insect Mol. Biol. 6, 183–190 22 Nogge, G. (1976) Sterility in tsetse flies (Glossina morsitans Westwood) caused by loss of symbionts. Experientia 32, 995–996 23 Southwood, T.R. et al. (1975) The micro-organisms of tsetse flies. Acta Trop. 32, 259–266

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