Chapter 3 The Enemy Within

The Enemy Within: Interactions Between Tsetse, Trypanosomes and Symbionts Deirdre P. Walshe, Cher Pheng Ooi, Michael J. Lehane and Lee R. Haines Liverpool School of Tropical Medicine, Pembroke Place, Liverpool L3 5QA, United Kingdom

1 Background 120 1.1 Human and animal trypanosomiases 120 1.2 Trypanosome species 121 1.3 Tsetse identification and distribution 122 1.4 Tsetse life cycle and physiology 123 1.5 Trypanosome (T. brucei sspp.) life cycle: Development and differentiation 126 2 Tsetse–trypanosome interactions 129 2.1 Parasite surface coat 130 2.2 Host blood factors 131 2.3 Tsetse midgut environment and signals for differentiation 133 2.4 Trypanosomes and tsetse digestive enzymes 134 2.5 Tsetse immune system 135 2.6 Effects of trypanosome infection on tsetse physiology 148 2.7 Fly sex, age and starvation and trypanosome transmission 149 2.8 Environmental temperature and trypanosome transmission 151 3 Symbiont–tsetse–trypanosome interactions 152 3.1 Wigglesworthia glossinidius 152 3.2 Wolbachia pipientis 153 3.3 Sodalis glossinidius 153 4 Towards new methods of disease control 156 4.1 Gene knockdown in Glossina 156 4.2 Paratransgenesis 158 5 Conclusion 159 Acknowledgements 160 References 160

ADVANCES IN INSECT PHYSIOLOGY VOL. 37 ISBN 978-0-12-374829-4 DOI: 10.1016/S0065-2806(09)37003-4

Copyright # 2009 by Elsevier Ltd All rights of reproduction in any form reserved

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Background HUMAN AND ANIMAL TRYPANOSOMIASES

African trypanosomiasis refers to a set of diseases of humans and their domesticated animals, which have devastating consequences for Africa. Tsetse flies (Diptera: Glossinidae) are the sole insect vectors responsible for cyclical transmission of African trypanosomes, the protozoan parasites responsible for Human African Trypanosomiasis (HAT ¼ sleeping sickness) and African Animal Trypanosomiasis (AAT ¼ nagana). In the early part of the last century several HAT epidemics occurred on the African continent, but by the early 1960s the disease was controlled and had almost disappeared (Steverding, 2008). However, relaxation of surveillance and control measures led to resurgence in HAT, peaking in 1997 at an estimated 450,000 cases (Barrett, 2006) with an estimated 60 million people at risk in 37 countries of sub-Saharan Africa (corresponding to one third of Africa’s total land area) (WHO, 2000). Since that date, case detection and treatment have been increased, and by 2007, the reported number of new HAT cases had dropped to 10,769 (WHO, 2007), which probably equates to 50–70,000 total human cases. HAT can take two forms depending on the parasite involved. Trypanosoma brucei rhodesiense and T. b. gambiense are the causative agents of HAT in East/Southern Africa and Central/West Africa, respectively. Typically, the T. b. rhodesiense transmission cycle involves wild and domestic animals, but intensified human to human transmission may occur during epidemics. The T. b. gambiense transmission cycle is mostly from human to human, involving animals to a much lesser extent. In humans, T. b. rhodesiense infections are acute, lasting from a few weeks to several months, while T. b. gambiense infections are chronic, generally lasting for several years, often without any major signs or symptoms. There are no prophylactic drugs or vaccines available to prevent HAT. In both cases, without proper diagnosis and treatment, the outcome is fatal. Therefore, the earlier the disease is detected the better the chance of survival. However, all four drugs currently used to treat HAT exhibit toxicity and, in many countries, drug resistance is beginning to emerge (Legros et al., 2002; Delespaux and de Koning, 2007; Balasegaram et al., 2009). Other parasite species and sub-species of the Trypanosoma genus are pathogenic to many wild and domestic animal species. In particular, T. b. brucei, T. congolense and T. vivax are major causes of the animal form of trypanosomiasis. Disease severity is dependent on both the pathogenicity of the parasite strain and the genetics of the mammalian host (Courtin et al., 2008). While most African wildlife is tolerant of the parasites, domesticated livestock are highly susceptible to disease, particularly if they originate from European stock. Despite implementation of many tsetse control strategies,

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nagana still has a large impact on agricultural and livestock production systems and land use. The disease has an estimated annual economic cost of approximately US $4.5 billion to the African economy due to losses in milk, meat and wool yields through adult mortality, calf mortality and subsequent depressed herd growth (Kristjanson et al., 1999). Furthermore, nagana is a major restriction to the development of arable agricultural in sub-Saharan Africa, limiting the use of draught and pack animals and preventing the development of mixed agricultural practices (Jordan, 1986). Currently, nagana is managed predominantly by use of trypanotolerant breeds of cattle and use of chemoprophylactic and trypanocidal drugs (Miruk et al., 2008).

1.2

TRYPANOSOME SPECIES

Two distinct groups of insect-transmitted trypanosomes are generally recognized. The Stercocaria (subgenus Megatrypanum, Schizotrypanum and Herpetosoma) and the Salivaria (subgenus Nannomonas, Duttonella and Trypanozoon). Stercorarian trypanosomes develop in the hindgut of the insect and are transmitted in the faeces. The predominant vectors of stercorarian trypanosomes are tabanids, triatomines, leeches and ticks. It is the salivarian trypanosomes which cause nagana and African sleeping sickness. The salivarian trypanosomes develop in the anterior part of the tsetse fly alimentary canal and are transmitted via the mouthparts. Tsetse flies are the major vectors of T. brucei trypanosomes but mechanical transmission of several salivarian trypanosome species, by tabanids and Stomoxys vectors, also occurs. Only the salivarian trypanosomes exhibit antigenic variation, a unique form of parasite immune evasion within the mammalian host (Cross, 1996) and it is several of these species which are responsible for the complex of diseases known as African trypanosomiasis. T. vivax (subgenus Duttonella), believed to be the most ancient of the salivarian trypanosomes, produce a high incidence of infection in the tsetse proboscis (Haag et al., 1998) and are serious pathogens of cattle. The Nannomonas subgenus is comprised of three species: T. congolense (Broden, 1904), T. simiae (Bruce et al., 1912) and T. godfreyi (McNamara et al., 1994). Of these, T. congolense is the most economically important, with a broad host range and wide geographical distribution. T. simiae and T. godfreyi are primarily associated with suid infections; the former being extremely pathogenic (acute) and the latter producing a chronic, sometimes fatal, infection in pigs. The Trypanozoon subgenus contains the trypanosomes causing HAT (T. b. gambiense and T. b. rhodesiense) and T. b. brucei which is one of the parasites causing nagana. T. b. brucei is unable to infect humans as it is sensitive to a trypanolytic factor in human serum (Oli et al., 2006) and is thus restricted to development in domestic animals and many species of wildlife.

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Of the African trypanosomes, T. congolense, T. b. brucei, T. b. gambiense and T. vivax have been selected for partial or complete genome sequencing by a consortium of institutes: The Institute for Genome Research (TIGR), now the J. Craig Venter Institute (Rockville, MD), and the Wellcome Trust Sanger Institute (Hinxton, UK). All genomes are in various states of completion. At the time of writing, the nuclear genome of T. brucei (TREU927 GUTat 10.1), the model species most often used for studying trypanosome biology, had been completed and the T. b. gambiense (MHOM/CI/86/DAL972) genome is in the last stages of finishing, with 8 coverage. Also, the partial nuclear genome sequence (5 coverage) of T. congolense (IL3000) and T. vivax are being assembled (http://www.sanger.ac.uk/Projects/Protozoa/).

1.3

TSETSE IDENTIFICATION AND DISTRIBUTION

The name tsetse (pronounced tsee–tsee) is derived from the noise the fly creates when it raises its body temperature by contraction of flight muscles decoupled from the wings, prior to energetically demanding events (e.g. birth of the larva, flight, rapid dehydration of the bloodmeal). Interestingly, tsetse means ‘‘fly’’ in the Tswana language and in Sechuana it is interpreted as ‘‘fly destructive to cattle’’. Tsetse flies are easily distinguishable from other insects (Fig. 1). They are light brown to black in colour and, dependant on the species, are roughly twice the size of a housefly. Characteristic aristae are present on the third antennal segment (Fig. 1A). In addition, the unique ‘‘hatchet’’ wing cell is found in the centre of each wing between the fourth and fifth veins (Fig. 1B). Also, tsetse flies adopt a characteristic resting attitude with their single pair of wings folded scissor-like over the dorsal surface of the abdomen. A

B

FIG. 1 Characteristic anatomical features of Glossina sspp. (A) Side view of an engorged female Glossina morsitans morsitans resting during diuresis. An anal droplet has started to form within minutes of feeding. The characteristic arista (arrow), a thin structure bearing a single unidirectional row of branched setae, is used for species identification. Photo: R. Wilson http://www.raywilsonbirdphotography.co.uk. (B) The middle of the wing contains a unique venation pattern resembling a hatchet, which is also characteristic of tsetse flies. Photo: L. Rafuse Haines.

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Tsetse flies fall into a single genus, Glossina, and are restricted to subSaharan Africa except for two localities in the Arabian peninsula. Twentythree species and eight sub-species of tsetse fly are currently recognized (Leak, 1999; Krafsur, 2009). These are divided into three distinct clades: Morsitans, Palpalis and Fusca, which are named after the best known species in each subgenus. Commonly these groups are described by the ecological niches they occupy; savanna (Morsitans), riverine (Palpalis) or forest (Fusca) groups. In Central and West Africa, the riverine species (Palpalis group) tend to feed predominantly on reptiles and ungulates. Humans regularly encounter these flies, particularly when visiting water sources, and these species are important vectors of human sleeping sickness. The savannah-woodlands species (Morsitans group) are the most economically important, as they preferentially feed on livestock and wildlife and are the major vectors of nagana. Both the Palpalis and Morsitans groups are vectors of T. brucei sspp. Most tsetse from the third clade (Fusca group) inhabit the damp, evergreen forests. The exception is G. brevipalpis, which is more regularly found in association with livestock. With the exception of G. brevipalpis, flies in the Fusca group are not considered to be medically or agriculturally important. Whether this will change when further pressure on land use drives them into more regular encounters with humans and their domesticated animals remains to be seen.

1.4

TSETSE LIFE CYCLE AND PHYSIOLOGY

Tsetse are unique among insect disease vectors in that they possess a viviparous lifestyle. Consequently, flies have a very low rate of reproduction, typically producing 8–10 offspring in their lifetime in optimal laboratory conditions (Leak, 1999; Attardo et al., 2006). The tsetse reproductive tract possesses extensive modifications to permit intrauterine larval development, which include a reduced number of ovarioles per ovary (two), a highly tracheated and muscular uterus and a modified uterine accessory gland (milk gland) to supply nutrients to the developing larvae (Leak, 1999; Attardo et al., 2006). Oogenesis begins before tsetse eclosion. A single oocyte develops at a time, starting with one of the two ovarioles in the right ovary, and takes 6–7 days to complete. Oogenesis appears to be regulated by the presence of a developing embryo or larvae in the uterus. In most Glossina species, the female is sexually mature 48–72 h posteclosion (i.e. emergence of the adult insect from the puparium), while males become fertile several days after eclosion. Female flies generally mate only once and can store sperm for the duration of their life. Upon completion of oogenesis, an oocyte is synchronously ovulated and fertilized in the uterus where it undergoes embryonic and larval development. The larva is solely nourished by a milk-like secretion rich in proteins and lipids produced by a pair of milk glands. A fully developed third instar larva is deposited by the

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female approximately 16 days after fertilization in the case of the first offspring, and every 9 days thereafter. The larva burrows into the ground where rapid pupariation takes place, and the adult fly emerges from the puparium approximately 4 weeks later. In the tsetse field, the term teneral is used to describe the period of time following emergence of the adult fly from the puparium until it takes its first bloodmeal. Both male and female adult tsetse flies are obligate haematophages capable of transmitting trypanosomes. Tsetse flies are pool feeders and the repeated penetration of mammalian host tissue by the tsetse proboscis results in the formation of a sub-surface blood pool. Saliva is expressed into the wound and trypanosomes are transmitted to the mammalian host at this stage. The bloodmeal is sucked up through the proboscis and oesophagus and propelled into the rest of the alimentary canal by the rhythmic pumping of the cibarial pump aided by the contraction of circular muscles that encompass the oesophagus. The proventriculus (¼cardia) (Fig. 2C) lies at the junction of the oesophagus, midgut and crop duct. Blood may pass directly into the midgut or into the extension of the foregut known as the crop (Fig. 2A), before regurgitation into the midgut (Moloo and Kutuza, 1970). The proventriculus acts as a valve regulating the directional flow of blood and is also the organ responsible for producing the peritrophic matrix (PM) (see Section 2.5.1). The tsetse midgut is a simple tube, lacking diverticula, running from the proventriculus to the junction with the hindgut, which is marked by the entrance of the Malpighian tubules into the alimentary canal (Fig. 2, top panel and Fig. 2J). Although more complex divisions exist (Bo¨hringer-Schweizer, 1977), the midgut can be crudely separated into three functional regions: the anterior midgut, bacteriome (¼mycetome) and posterior midgut. The first part of the anterior midgut is a linear tube running though the thorax. Once it enters the abdomen, the anterior midgut becomes distended and here the blood is stored and dehydrated prior to digestion. Epithelial cells of the anterior midgut possess extensive infoldings of the basal plasma membrane with associated mitochondria, enabling the fly to achieve this rapid dehydration (Bo¨hringerSchweizer, 1977). The anterior midgut is interrupted approximately in its middle section by a region of cells called the bacteriome (Fig. 2E). The bacteriome houses the intracellular symbiotic bacteria Wigglesworthia glossinidius. Haemolysis and bloodmeal digestion commence at the very beginning of the posterior midgut, where haemolytic agents and digestive enzymes are produced. Virtually all proteolytic enzymes are restricted to the posterior midgut (Gooding, 1974a). The junction of the anterior and posterior midguts (Fig. 2F) is obvious and tightly delineated in fed flies, as the bloodmeal changes in colour from red to brown/black as haemolysis and digestion progesses. Cells of the proximal part of the posterior midgut possess extensive rough endoplasmic reticulum, large numbers of Golgi bodies and secretory vesicles allowing efficient production, storage and secretion of proteins (Bo¨hringer-Schweizer, 1977).

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A Posterior Midgut

Anterior Midgut

F B

Hindgut

I

C

J K

D E

L

G H

A

B

C

D

E

F

G

H

I

J

K

L

FIG. 2 Anatomy of the alimentary canal of Glossina morsitans morsitans. Top panel represents a dissected alimentary canal aligned anterior (left) to posterior (right). The letters (A–L) correspond to the lower panels, which are magnified regions of the digestive system visualized using light microscopy; (A) the crop, (B) salivary glands, (C) the proventriculus capped by a modified fat body crown (oenocytes), (C–D) the most proximal part of the anterior midgut that lies in the thorax, (D) the point where the anterior midgut enters the abdomen, (E) the bacteriome that contains the obligate symbiont Wigglesworthia glossinidia, (F–G) the most distal part of the anterior midgut and the junction where the anterior and posterior midgut meet, demarked by a bloodmeal colour change (from red to black), (H) fat body attached to the wall of the posterior midgut, (I) the distinctive posterior midgut epithelial cells (luminal surface), (J) the bases of the Malpighian tubules leave the alimentary canal at the junction of the midgut and hindgut. The two Malpighian tubules each branch near their base to form a total of four tubules, (K) ileum (severed from the MT and midgut) and the junction with the colon, (L) colon ending in the bulbous rectum; note peritrophic matrix (*) exuding from a breach in the hindgut wall. Photo: L. Rafuse Haines.

Cells of the distal part of the posterior midgut are involved in absorption of digested products. They possess extensive smooth endoplasmic reticulum and also lipid droplets and glycogen at various times of the digestive cycle (Bo¨hringer-Schweizer, 1977).

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DEIRDRE P. WALSHE ET AL. TRYPANOSOME (T. BRUCEI SSPP.) LIFE CYCLE: DEVELOPMENT AND DIFFERENTIATION

T. brucei has the most complex, but perhaps the best characterized, life cycle of all African trypanosome species. The trypanosome life cycle was first described in detail by Muriel Robertson who described the successive stages of parasite establishment and maturation within the insect and mammalian hosts, demonstrated the migration of parasites through the fly midgut and proved that only salivary gland forms were capable of producing a mammalian infection (Robertson, 1913). Since then, a more complete understanding of trypanosome development has been achieved, with an agreed parasite nomenclature adopted (Roditi and Clayton, 1999) and a consensus achieved on many of the barriers present in the fly that the trypanosome must overcome to survive and develop in order to complete cyclical transmission. Within the vertebrate bloodstream at least two different major forms of trypanosomes are found; a long slender form, which replicates by asexual division, and a short stumpy, non-replicating form (Fig. 3(1)). These extracellular parasites are covered with an immunogenic surface coat composed of approximately 107 identical variant surface glycoprotein (VSG) molecules (Vickerman, 1969; Barry and McCulloch, 2001; Barry et al., 2005). The VSG coat physically shields underlying membrane proteins from host immune responses and is central to antigenic variation and survival in the mammalian host. The consecutive, but mainly unpredictable, expression of a large repertoire of VSG genes permits expansion of antigenically distinct trypanosome populations within the host. After activation of host immune responses (in reaction to high parasitaemia), the majority of the parasite population is destroyed. A small number of trypanosomes survive because they express an antigenically distinct VSG coat, and proceed to expand in numbers. The continuous cycles of trypanosome replication and destruction result in waves of fluctuating parasitaemia. The differentiation of the long slender bloodstream form (BSF) into the non-dividing stumpy BSF occurs in high density populations of long slender BSFs (Vassella et al., 1997; Seed and Wenck, 2003). The switch to stumpy BSF involves changes in metabolism within the trypanosome, but the molecular signals involved are not yet known. Short stumpy BSFs are believed to be pre-adapted for survival within the insect midgut due to the presence of a functional mitochondrion. In the vertebrate bloodstream, trypanosomes utilize glucose as an energy source. However, in the fly midgut, glucose is limiting and a more efficient utilization of glucose and amino acids occurs via the Krebs cycle and oxidative phosphorylation in the mitochondrion. To study the early events of trypanosome establishment in the tsetse midgut, T. b. brucei trypanosomes expressing green fluorescent protein (GFP) under the control of a procyclin promoter have been created (Gibson and Bailey, 2003). It is evident that differentiation of the BSF to the procyclic form (i.e. the insect

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4. Proventriculus

LE

6. Salivary glands M

SE

ADT

SE Ms

LT

ADT

Peritrophic matrix Epithelium SS Ms

P

LS

1. Bloodmeal

3. Ectoperitrophic space P

SS

2. Endoperitrophic space

FIG. 3 Diagram of the life cycle stages of T. b. brucei within the tsetse fly. Trypanosome morphology was based on observations by Van den Abbeele et al. (1999) and Sharma et al. (2008). Stages of the trypanosome life cycle within the tsetse are indicated within circles joined by the red arrow; the grey arrow represents transit between insect and mammalian host. 1. Short stumpy (SS) and long slender (LS) bloodstream forms (BSFs) ingested with the infective bloodmeal; 2. Transformation of short stumpy (SS) BSF into procyclics (P) within the midgut endoperitrophic space; 3. Procyclics differentiate into mesocyclics (Ms) within the ectoperitrophic space; 4. Transformation of mesocyclics (Ms) into long trypomastigotes (LT), which give rise to asymmetrically dividing trypomastigotes (ADT); 5. Trypomastigotes (ADT) divide into long epimastigotes (LE) and short epimastigotes (SE). These three stages are present in both the proventriculus and salivary glands; 6. Short epimastigotes (SE) interdigitate within the salivary gland epithelium at their anterior ends, and subsequently mature into the nondividing, mammalian infective metacyclics (M). Note the gradual migration of the kinetoplast in the fly stages of the parasite, culminating in a position anterior to the nucleus in the trypomastigote and epimastigote forms. The reversion of this position to the posterior end of the parasite is a signature of mature metacyclics.

midgut-adapted form), which involves replacement of surface VSGs by procyclins, occurs rapidly after bloodmeal ingestion by the fly (Fig. 3(2)) (Vassella et al., 2001; Acosta-Serrano et al., 2001; Gibson and Bailey, 2003). Most flies successfully kill all invading trypanosomes in a process termed self-cure.

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For the first 3 days, trypanosomes are mostly contained within the bloodmeal as it is being digested. The critical events in parasite establishment appear to occur approximately 3 days after infection, when the relatively small proportion of surviving trypanosomes ( 10%) either die or rapidly multiply in number (Gibson and Bailey, 2003). Typically, from 8 days after the infected bloodmeal, dissected flies can be confidently divided into two groups; the first in which most flies will have self-cured (having completed the clearing of ingested trypanosomes from their midguts) and the second which have established midgut infections. Trypanosomes in an established infection migrate to the ectoperitrophic space 3–5 days post-infection (Gibson and Bailey, 2003) (Fig. 3(3)). It is believed that this occurs by direct penetration through the PM (Ellis and Evans, 1977; Gibson and Bailey, 2003) although an alternative but less likely, suggestion is that it occurs by circumnavigation around the open, posterior end of the PM in the hindgut (see Section 2.5.1). Typically the midgut population in an established infection reaches approximately 5  105 trypanosomes (Van den Abbeele et al., 1999; Gibson and Bailey, 2003). From 6 to 8 days post-infection, large numbers of trypanosomes congregate within the proventriculus (Van den Abbeele et al., 1999; Gibson and Bailey, 2003; Sharma et al., 2008) (Fig. 3(4)). Here they appear to cease division, elongate to mesocyclic forms and later differentiate into long trypomastigotes (Fig. 3(4)) (Van den Abbeele et al., 1999). Trypanosomes then migrate back into the endoperitrophic space by actively penetrating the PM and move anteriorly in the lumen of the foregut to the opening of the hypopharynx at the tip of the proboscis. An alternative theory of migration involves the direct penetration of the tsetse salivary glands after trypanosomes have traversed the fly haemolymph (Mshelbwala, 1972). It is generally accepted that this is unlikely, as trypanocidal factors known to be present in the haemolymph (Croft et al., 1982) would act as a major barrier for trypanosomes attempting to traverse it. Early positioning of trypanosomes in the anterior midgut and proventriculus should also favour passage along the foregut to the salivary glands (Peacock et al., 2007). Asymmetric division of the proventricular epimastigote form generates both long and short parasites (Fig. 3(5)) and it is either the asymmetrically dividing trypanosome or the short epimastigote that arrives at the salivary gland (Sharma et al., 2008). Each tsetse fly has two salivary glands. Evidence suggests that each gland is invaded and colonized separately, with few epimastigotes constituting the founder populations (Peacock et al., 2007). The short epimastigote forms are believed to attach to the salivary gland epithelium by interdigitation of their membranes (Fig. 3(6)). Upon binding, the non-infective epimastigotes complete several rounds of replication and differentiate into the metacyclic form. Differentiation (metacyclogenesis) includes the appearance of a VSG surface coat. Metacyclic VSGs display a specific VSG repertoire subset and their expression is regulated differently to bloodstream VSGs (Barry et al., 1998; Graham et al., 1999). Mitochondrial changes also occur, including loss of mitochondrial cristae and Krebs cycle enzymes. The biochemical changes

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accompany the posterior migration of the kinetoplast before the parasite detaches into the lumen as a mature, free-form, infective metacyclic trypomastigote. At this point, each mature metacyclic parasite has undergone the transformations necessary for survival in a mammalian host. The above is an account of the most complex trypanosome life cycle completed in the tsetse fly and it should be noted that there are distinct differences between the life cycle of T. brucei sp. trypanosomes and other African trypanosome species. T. vivax completes its entire life cycle in the proboscis of the fly while T. congolense infections develop in the fly midgut and mature in the hypopharynx. The number of flies that develop a mature infection and the length of time required for an infection to establish and mature into a transmissible form can vary depending on several factors, including fly species (Welburn et al., 1989, 1994), fly sex (Distelmans et al., 1982; Dale et al., 1995) (see Section 2.7) and parasite strain (Dale et al., 1995). Recent evidence suggests migration of trypanosomes from the proventriculus to the salivary glands may occur continuously (Peacock et al., 2007; Sharma et al., 2008), albeit at low numbers, as opposed to being restricted to a limited time ‘‘window’’ (Van den Abbeele et al., 1999). Once a fly is infected it will produce infective metacyclics for the duration of its life, which can be 150 or more days for females and about half that for males (Lehane and Mail, 1985; Msangi et al., 1998). Thus, there is potential for infective parasites to be transmitted every time a fly feeds on a new host. In addition, flies are capable of harbouring mixed infections of two or more species of trypanosome (Lehane et al., 2000; Van den Bossche et al., 2004; Kubi et al., 2005; Peacock et al., 2007). Sexual reproduction has been reported in T. b. brucei in the salivary glands and may exist in other trypanosome species (Jenni et al., 1986; Peacock et al., 2007; Gibson et al., 2008). However, mating is not an obligatory part of the trypanosome life cycle and occurs in only a proportion of flies co-infected with two different strains of T. b. brucei (Jenni et al., 1986; Schweizer et al., 1988). It is suggested that the unattached epimastigote is the mating stage (Gibson et al., 2008) (Fig. 3) but the mechanism of genetic exchange is not yet known. Under field conditions mating may be a rare event (Koffi et al., 2009).

2

Tsetse–trypanosome interactions

It appears from the established literature that all tsetse species are susceptible, to some degree at least, to trypanosome infections. In general, tsetse in the Palpalis group species tend to be poor vectors of congolense-type trypanosomes compared to the Morsitans group flies (Harley and Wilson, 1968; Moloo and Kutuza, 1988a; Ndegwa et al., 1992). Conversely, tsetse of the Morsitans group are poorer vectors of T. b. gambiense than the Palpalis group (Richner

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et al., 1988). Care needs to be taken with much of the data on susceptibility, as often fly and trypanosome strains used in experiments are from widely divergent geographical origins. Many factors influence fly susceptibility to trypanosome infection. Our understanding of these factors and their underlying mechanisms is still rudimentary. In this review, we concentrate on the physiological factors influencing tsetse fly susceptibility to trypanosomes. 2.1

PARASITE SURFACE COAT

Throughout their life cycle in the tsetse fly, trypanosomes are covered by different surface molecules. Successful differentiation of BSF to procyclic forms involves the shedding of surface VSG and replacement with a set of new surface molecules, the best known being the procyclins (Roditi and Pearson, 1990; Beecroft et al., 1993). The procyclins are major GPI-anchored proteins possessing extensive C-terminal glu-pro (EP) or gly-pro-glu-glu-thr (GPEET) repeats, and comprise a surface coat of approximately three million procyclin molecules per cell (Roditi et al., 1998; Pays and Nolan, 1998; Roditi and Lehane, 2008). Three isoforms of EP procyclin, EP1, EP2 and EP3, are known, which differ in the length of their repeats, the sequence of their N-terminal domains and the presence or absence of N-glycosylation sites. Analysis of procyclin expression by mass spectrometry indicated that trypanosomes exhibit distinct procyclin expression profiles in vivo. All four procyclins are present at similar levels within a few hours of differentiation to procyclics (Vassella et al., 2001). However, 3 days post-infection, the procyclic coat consisted primarily of GPEET with lower amounts of EP forms (AcostaSerrano et al., 2001). This GPEET was 11 residues shorter than that of in vitro cultured procyclic culture forms (PCFs). From 7 days post-infection, GPEET procyclin disappeared and expression switched to the glycosylated isoforms EP1 and EP3. EP1 and EP3 were also truncated in comparison to these isoforms expressed by in vitro cultured PCF. Transcripts of EP2 are quite abundant in fly-derived procyclic forms but EP2 proteins have not yet been confirmed in vivo (Urwyler et al., 2005). While tsetse procyclins exhibit complex expression profiles (suggesting a selective advantage in their production) their function remain unknown. Proteolysis in the fly removes the N-terminal domains of all procyclins, but the acidic amino acid repeat sequences are largely resistant to proteolysis (Acosta-Serrano et al., 2001). Thus procyclins may serve a protective role, aid parasite development and/or influence ligand-associated parasite–vector signalling (Roditi and Pearson, 1990; Ruepp et al., 1997). Interestingly, parasites expressing procyclins with truncated N termini can still establish midgut infections to similar levels as wild-type parasites under laboratory conditions and form mature metacyclics within the salivary glands (Liniger et al., 2004). Thus, procyclins are not crucial for migration of procyclic forms from the tsetse midgut to the salivary glands (Vassella et al., 2009). However, in co-infection experiments,

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procyclin null mutants were rapidly outgrown in the midgut by wild-type parasites. This suggests that under field conditions, where mixed infection in flies is common (Lehane et al., 2000), surface procyclins probably play a significant role in trypanosome fitness (Vassella et al., 2009). As trypanosome epimastigotes within the tsetse salivary glands typically lack a procyclin coat (Urwyler et al., 2005), the function of the procyclin coat molecules probably occurs earlier in trypanosome development in the fly. T. congolense PCFs express several major stage-specific surface molecules different to those of T. brucei. In the early stage of T. congolense midgut infection, a protease-resistant surface molecule (PRS) is expressed, while later, a heptapeptide-repeat containing molecule (Butikofer et al., 2002) and a glutamic acid–alanine-rich protein (GARP) are expressed (Beecroft et al., 1993; Bayne et al., 1993; Utz et al., 2006). Interestingly, T. congolense procyclins bear similarity to T. brucei procyclins in being acidic and having repeat sequences rich in glutamic acid, glycine, threonine and proline (Utz et al., 2006). Also, genes related to GARP have been identified in both T. simiae and T. godfreyi. Epimastigote forms of T. congolense and T. brucei, found in the proboscis and salivary glands respectively, express related surface glycoproteins named congolense epimastigote specific protein (CESP) (Inoue et al., 2000) and brucei alanine-rich proteins (BARP) (Urwyler et al., 2005). The function of these different parasite surface molecules is still unknown. In the salivary glands nascent and mature metacyclics (re)acquire a VSG coat in preparation for transfer to the next mammalian host (Tetley and Vickerman, 1985). 2.2

HOST BLOOD FACTORS

Different trypanosome species are capable of infecting different mammalian hosts, which implies that host blood factors can affect trypanosome survival (Vickerman, 1985; Masaninga and Mihok, 1999). An intriguing question is whether or not the influence of these host blood factors extends to trypanosome establishment in the tsetse midgut. T. brucei developmental forms are present in the midgut for at least 8 days, before commencing migration and maturation in the foregut and salivary glands. As tsetse flies ingest a new bloodmeal every 24–48 h, there is ample opportunity for an ingested bloodmeal to have an impact on trypanosome survival. The mammalian host blood in which the trypanosome infection is acquired by tsetse affects both differentiation (Nguu et al., 1996) and multiplication of parasites in the fly midgut (Olubayo et al., 1994; Mihok et al., 1995). Buffalo and eland blood support establishment of T. congolense infections in G. m. morsitans less well than goat blood (Olubayo et al., 1994). This was apparent from 3 days after the infective bloodmeal and occurred regardless of whether flies were maintained on rabbit blood or the blood of the respective mammalian hosts in which the infective meal was given (Olubayo et al., 1994).

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This suggests that the trypanocidal event in the fly midgut occurs early in the infection process. Serum components of the blood may be either conducive or detrimental to survival of trypanosomes in the fly (Reduth et al., 1994; Mihok et al., 1995; Muranjan et al., 1997). The serum of the Cape buffalo, for example, is trypanocidal to most species of trypanosomes (Reduth et al., 1994). Using T. b. brucei BSF, the primary cause of parasite death was inhibition of glycolysis by host hydrogen peroxide (H2O2) generated by xanthine oxidase (Muranjan et al., 1997). Whether or not this effect holds true for procyclic forms if a recently infected fly subsequently feeds on Cape buffalo merits a more detailed study. However, it is unlikely the same trypanocidal mechanism would occur in vivo since procyclic trypanosomes metabolize proline, rather than glucose, as a major energy source (Muranjan et al., 1997). While xanthine oxidase is present in the serum of other mammals, it does not cause death of different trypanosome species in these hosts (Cruz et al., 1983; Reduth et al., 1994). Therefore, it is possible that generation of H2O2 to trypanocidal levels by xanthine oxidase in Cape buffalo may be part of a wider metabolic cascade, including purine catabolizing enzymes (Muranjan et al., 1997). Thus, experiments using the serum of different mammalian species, which show seemingly little correlation between serum concentration and BSF trypanosome survival, may simply differ in the molecular constituents of different mammalian host serums (Black et al., 1999). Host serum complement (an immunological cascade present in mammalian serum, which, upon activation, induces lysis of target microbes) may also be involved in causing trypanosome mortality (Ferrante and Allison, 1983; Black et al., 1999). A BSF trypanocidal factor, which is heat labile (inactivated following 30 min at 56  C), has been documented in the serum of mammalian species lacking any xanthine oxidase activity (Black et al., 1999). Also, T. congolense and T. b. brucei PCFs are killed by another heat sensitive factor in human serum (Ferrante and Allison, 1983). Similar PCF lytic activity was observed in human serum deficient in complement C2 (part of the classical complement pathway), as well as in ethylene glycol tetra-acetic acid (a preferential chelator of calcium ions, a crucial substrate for initiation of the classical pathway). Consequently, the trypanocidal factor may rely on the alternative complement pathway for activation (Ferrante and Allison, 1983). The infective bloodmeal origin also appears to play a key role in determining the virulence and infectivity of the resulting salivary gland metacyclics (Masaninga and Mihok, 1999). In vitro grown T. congolense BSFs were mixed with either goat, eland or zebra blood and fed to G. m. centralis flies. The resulting metacyclics used to infect BALB/c mice varied in their prepatent periods and resulted in differences in mouse mortality rates (Masaninga and Mihok, 1999). A strong selection pressure appears to be exerted by host blood, most likely on the procyclic parasite stage exposed to incoming bloodmeals, which may lead to differential killing of parasites.

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133

TSETSE MIDGUT ENVIRONMENT AND SIGNALS FOR DIFFERENTIATION

Efforts to determine the triggers responsible for trypanosome switching from bloodstream to procyclic forms have primarily been conducted in vitro and have occasionally ignored the realities of tsetse fly biology. A variety of methods have been used to define differentiation including morphological markers (Hunt et al., 1994), the release of VSG proteins and concomitant detection of procyclins, or the detection of DNA synthesis (Matthews and Gull, 1997). Other novel approaches include the use of transgenic trypanosomes carrying the E. coli glucuronidase (GUS) gene under the control of procyclin expression elements (Sbicego et al., 1999) and the use of GFP-expressing trypanosomes under the regulation of a procyclin-linked promoter (Sheader et al., 2004). Reduction of temperature alone from 37 to 27  C can initiate the differentiation of BSFs to PCFs (Brown et al., 1973; Bienen et al., 1980; Overath et al., 1986) and is conducive to growth and proliferation of a transformed trypanosome population. Using the GUS system, Sbicego et al. (1999) found that both citrate and cis-aconitate (Krebs cycle intermediates) could induce trypanosome differentiation to PCF in vitro, in a manner independent of the commonly used temperature shift (from 37 to 27  C). This phenomenon appears to be highly specific as other Krebs cycle intermediates, even when present at relatively high concentrations in vitro (10 mM), did not trigger parasite differentiation (Sbicego et al., 1999). Also, compounds closely resembling citrate and cis-aconitate, for example trans-aconitate and 5-fluoro-citrate, were trypanocidal (Hunt et al., 1994). To date, the citrate concentration of the fly midgut lumen has not been reported. However, if similar to the low citrate concentration estimated within the fly haemolymph (15.9 mM), this may be insufficient to trigger BSF differentiation (Hunt et al., 1994). However, BSFs exhibit hypersensitivity towards cis-aconitate when they are exposed to a cold shock (Engstler and Boshart, 2004). Thus lower concentrations of citrate and cis-aconitate may be sufficient to trigger differentiation of BSFs in vivo when used in combination with a temperature drop (Hunt et al., 1994; Rolin et al., 1998; Fenn and Matthews, 2007). Trypanosome differentiation may also be initiated in vitro by growth of BSFs in low glucose medium (Milne et al., 1998). This may reflect what occurs in vivo, as changes in glucose concentration occur as the trypanosomes transfer from a high glucose environment (mammalian bloodstream) to a lower glucose environment (fly midgut). Furthermore, glucose concentration may be a factor involved in controlling switches in procyclin expression at later stages of trypanosome development (Morris et al., 2002). Incubation of BSFs under mildly acidic conditions (pH 5.5) was shown to cause differentiation to PCFs in vitro (with corresponding shedding of the VSG coat and expression of procyclins) at 27  C, even in the absence of cis-aconitate (Rolin et al., 1998). However, this in vitro work seems to have ignored conditions in the tsetse fly, where the best available data suggest that the pH of the midgut is highly alkaline. The use of microelectrodes to measure the midgut pH

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in unfed flies and fed flies 48 and 72 h post-feed has shown that the tsetse digestive tract average pH lies between 9 and 9.5, with the lowest pH value (pH 7.9) occurring in the posterior midgut 72 h post-feeding (Liniger et al., 2003). The enzymes trypsin (see Section 2.4) and thermolysin stimulate trypanosome differentiation in vitro. In the case of trypsin, at least, this occurs independently of any slender BSF attrition effects (Sbicego et al., 1999). Pronase was also shown to be an efficient trypanosome differentiation trigger, with up to 95% differentiation to PCFs achieved (Hunt et al., 1994). However, as far as we are aware, of the proteases above only trypsin is found in the tsetse midgut (Gooding and Rolseth, 1976). Another molecular trigger native to the tsetse midgut, the Glossina proteolytic lectin serine protease (Gpl) isolated from Glossina fuscipes fuscipes, has been identified as having a role in trypanosome differentiation in vitro (Abubakar et al., 2006). To date, while several triggers of differentiation from BSF to PCFs have been identified, the signals that cause differentiation to epimastigote and metacyclic forms in vivo are unknown. 2.4

TRYPANOSOMES AND TSETSE DIGESTIVE ENZYMES

Tsetse flies are obligate haematophages and thus are completely specialized for digesting blood. The majority of their nutritional resources are obtained from the protein content of the bloodmeal (Moloo, 1976; Kabayo and Langley, 1985). Consequently, the fly has a range of proteolytic digestive enzymes in the posterior, digestive portion of the midgut (Gooding and Rolseth, 1976; Cheeseman and Gooding, 1985). The anterior portion is virtually free of proteolytic activity and indeed the epithelium of the anterior midgut actively secretes proteinase inhibitors into the gut lumen (Gooding, 1974b; Houseman, 1980; Stiles et al., 1991). It has previously been suggested that midgut proteases are a major barrier to trypanosome establishment within the tsetse midgut (Imbuga et al., 1992). Given the above, while this may be the case, it probably only refers to trypanosomes entering the posterior midgut. In addition, fly midgut trypsin levels do not differ in infected and refractory flies, suggesting that digestive enzymes are not part of a protective response to trypanosome infection. Neither feeding excess trypsin nor inhibition of trypsin results in any difference in infection phenotype within the fly (Welburn and Maudlin, 1999). Studies using tsetse midgut extracts have found no correlation between tsetse gut protease activity and trypanosome infection rates (Mihok et al., 1994). Collectively these data indicate that it is questionable whether tsetse proteases have any major bearing on trypanosome infections within the fly gut. Curiously, an increase in mRNA transcript levels of certain digestion related enzymes, namely the tsetse cathepsin B, zinc carboxypeptidase and zinc metallo-protease, have been reported when a trypanosome infection is present (Yan et al., 2002). It should be noted however that protein levels, particularly in the intestine of insects taking occasional meals, may not be proportional to transcript levels.

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135

TSETSE IMMUNE SYSTEM

Despite their obvious efficiency in maintaining large burdens of trypanosomebased disease in Africa, tsetse flies exhibit a considerable level of refractoriness to trypanosome infection. Even under optimal laboratory conditions, where flies are fed at regular intervals, only a proportion of flies will establish midgut infections and the number decreases dramatically after the adult fly has taken three to four bloodmeals (Fig. 4) (Distelmans et al., 1982; Welburn and Maudlin, 1992; Kubi et al., 2006). Furthermore, less than half of the infections that become established in the midgut will mature (Van den Abbeele et al., 1999; Gibson and Bailey, 2003; Peacock et al., 2006). A key factor in this refractoriness is the fly immune system (Hao et al., 2001). Immune stimulation, by injection of live E. coli or lipopolysaccharide (LPS) into the haemocoel of the fly prior to feeding an infective bloodmeal, leads to a statistically significant decrease in trypanosome midgut infection rates (Hao et al., 2001). Identification of the tsetse immune molecule(s) responsible for conferring resistance to trypanosome infection has been hampered by the lack of an annotated Glossina genome (scheduled for completion in 2011). However, the sequencing and annotation of EST libraries from several tissue sources,

100

Trypanosome prevalence (%)

90 80 70 60 50 40 30 20 10 0

1st BM n = 307 r = 11

2nd BM n = 165 r=5

3rd BM n = 333 r = 12

4th BM n = 117 r=4

5th BM n = 100 r=2

8th BM n = 52 r=2

10th BM n = 35 r=1

FIG. 4 G. m. morsitans teneral phenomenon. The timing of the infective bloodmeal (BM) affects the prevalence of establishment of trypanosomes in the tsetse midgut. The infective BM containing T. b. brucei (TSW196) BSF given at one of the stated bloodmeals (x-axis) results in the indicated prevalence of infection (mean  SE). N ¼ cumulative sample size number (male), R ¼ number of replicates (Lehane laboratory, unpublished results).

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including the major immunoresponsive tissues of midgut (Lehane et al., 2003) and fat body (Attardo et al., 2006), has provided the foundation for more extensive studies of the Glossina innate immune system at a molecular level (http://www.genedb.org/genedb/glossina/index.jsp). Based largely on work on Drosophila melanogaster, it is known that insects possess a complex, interacting, innate immune system. This system is comprised of physical barriers (such as the cuticle and the PM), cellular responses (such as encapsulation and phagocytosis), and humoral responses, such as the generation of host defence peptides (HDP, previously called antimicrobial peptides), reactive oxygen species (ROS) and melanization by the phenoloxidase pathway (Lemaitre and Hoffmann, 2007). The immune response depends not only on the nature of the immune stimulus, but also its route of delivery, with quite distinct epithelial and systemic immune response profiles generated to the same pathogen (Hao et al., 2001). Clearly, in tsetse–trypanosome interactions, it is the epithelial immune responses of the alimentary canal and salivary gland tissues that are likely to be of major importance, as trypanosomes involved in the natural life cycle are exposed only to epithelial surfaces throughout the parasite life cycle. Trypanosomes have been reported in the tsetse haemocoel (Mshelbwala, 1972; Otieno et al., 1976), but these are almost certainly not important to the completion of the normal life cycle. Those trypanosomes that do traverse the midgut epithelium are rapidly killed by an unidentified systemic immune response (Croft et al., 1982), which effectively confines the trypanosomes to the lumen of the alimentary canal.

2.5.1

Peritrophic matrix

The insect midgut epithelium is physically protected from abrasion by food (and the pathogens it may contain) by the peritrophic matrix (PM, previously known as the peritrophic membrane) (Lehane, 1997). The PM is composed of a highly organized, glycosaminoglycan-rich layer reinforced with chitin (Tellam et al., 1999). In tsetse, the PM is constitutively expressed, forming a protective sleeve along the entire length of the midgut. Type II PM, such as that occurring in tsetse flies, physically separates the midgut lumen into two compartments. The endoperitrophic space is where the food lies and the ectoperitrophic space is the region between the PM and the midgut epithelium (see inset, Fig. 3). Clearly, by separating the food from the midgut epithelium, the PM must act as a molecular sieve through which digestive enzymes and nutrients destined to be absorbed must pass. The molecular sieving properties of the PM vary with changes in the surrounding ionic environment (Miller and Lehane, 1993). Typically, the tsetse PM has a pore size of 9 nm, making the PM permeable only to globular molecules less than 150 kDa (Miller and Lehane, 1990). Consequently, the tsetse PM presents a barrier to procyclic trypanosomes that need to get into the ectoperitrophic space to continue their development.

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Whether trypanosomes penetrate the PM, or circumnavigate around its broken ends in the hindgut to get to the ectoperitrophic space, is still the subject of speculation. There is little dispute that trypanosomes returning to the endoperitrophic space, from 6 to 8 days post-infection, actively penetrate the PM. Given the data obtained mainly by transmission electron microscopy (Ellis and Evans, 1977; Gibson and Bailey, 2003), it seems probable that trypanosomes also actively penetrate the PM when entering the ectoperitrophic space 3–5 days post-infection. Penetration is likely to be preceded by a specific attachment event, so it is interesting to note that GFP-expressing T. b. brucei PCFs tend to line up parallel with the PM and become associated with tsetse gut fragments upon dissection (Gibson and Bailey, 2003). How trypanosomes achieve PM penetration is unknown, and no chitinase gene has been identified from the trypanosome genome. However, chitin can be a relatively minor component of dipteran PM (Tellam et al., 1999) and other enzymes capable of dealing with glycosaminoglycans may be more important in the penetration event. In support of penetration rather than circumnavigation via the hindgut, only small numbers of procyclics were observed in the posterior region of the midgut when GFP-expressing parasites were used. These parasites were considered to be an indication of a failed infection cycle in its terminal stages (Gibson and Bailey, 2003).

2.5.2

Immune signalling

Genetic studies have demonstrated that Drosophila uses two main immune signalling pathways to control immune responses to pathogen challenge. The Toll and Imd (immunodeficiency) pathways respond to different classes of microbes and are used differently in various fly tissues (De Gregorio et al., 2002). The Imd pathway is predominantly involved in regulating epithelial immune responses and is most strongly utilized following Gram-negative bacterial challenge. The Toll pathway is most heavily involved in systemic immune responses and is activated most by fungal and Gram-positive bacterial infections (Fig. 5). These pathways are highly conserved among insect species and bear similarity to the vertebrate Toll-like receptor (TLR) and tumour necrosis factor (TNF) pathways. The Drosophila immune system is also highly regulated as over- or underexpression of immune responses is detrimental to host health (Lemaitre and Hoffmann, 2007; Ryu et al., 2008; Aggarwal and Silverman, 2008). The immune response may be divided into three key stages: recognition of pathogens, activation of the signal transduction pathway(s) and production of immune effector and regulator molecules. Orthologous intracellular and extracellular members of both pathways have been identified from Glossina EST databases (Table 1), although very little experimentation to gather functional evidence for their role in tsetse flies has been undertaken.

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Toll

Imd

Yeast

Fungi

Gram-positive bacteria

Gram-negative bacteria

GNBP3 Lysine-type peptidoglycan

DAP-type peptidoglycan PGRP-LE

Haemolymph

Necrotic GNBP1

PGRP-SD

Monomeric peptidoglycan

SPE

Persephone

Polymeric peptidoglycan

PGRP-SA Pro-Spätzle PGRP-LCx PGRP-LCa PGRP-LCx PGRP-LCx

Spätzle

PGRP-LF Imd

Toll

MYD88

dIAP2 Tube

Pelle

Caspar

dFADD

Cytoplasm

dTAB2 dTAK1

Dredd Dnr1

Cactus WntD

Dorsal

DIF

JNK pathway

Nucleus

DaPKC Drosomycin + other HDPs

Relish Kenny Ird5 (IKKγ) (IKKβ) Caudal Diptericin + other HDPs

FIG. 5 Schematic representation of the Toll and Imd signalling pathways. The function of the Toll and Imd signalling pathways has been intensively investigated in Drosophila melanogaster and has been reviewed in detail by Lemaitre and Hoffmann (2007) and Aggarwal and Silverman (2008). This figure is modified from these references. The omniBLAST search tool was used to identify putative G. m. morsitans orthologs (at 1e 20 or less) of all identified Drosophila Toll and Imd pathway genes in all library clustered EST translated sequences of G. m. morsitans (http://www.genedb.org). Orthologs of all Toll and Imd pathway members depicted were identified (except WntD) and are summarized in Table 1.

2.5.3

Recognition and activation

Differential regulation of immune effector HDP indicates that the tsetse immune system can distinguish between virulent and avirulent strains of bacteria (Weiss et al., 2008), between trypanosome and bacterial infections and also between different trypanosome life stages, that is procyclic and bloodstream forms (Hao et al., 2001). In Drosophila, pathogen recognition is achieved by pattern recognition receptors (PRRs), which each recognize a certain class of pathogen-associated molecular pattern (PAMP). Two families of PRRs, the peptidoglycan recognition proteins (PGRPs) and Gram-negative binding proteins (GNBPs), are known. Interestingly, PRRs are proteins originally derived from enzymes known to degrade microbial cell wall components, thus evolving

TABLE 1 Identification of Toll and Imd immune pathway orthologs in G. m. morsitans Toll Drosophila Molecule

Imd a

b

Ortholog

Drosophila Molecule

GNBP1 (Gram-negative FBgn0040323 binding protein 1) GNBP3 (Gram-negative FBgn0040321 binding protein 3) Persephone FBgn0030926

cn16500

Necrotic

FBgn0002930

cn8210

SPE (Spa¨tzle processing FBgn0039102 enzyme) Spa¨tzle FBgn0003495 Toll FBgn0003717

cn8446

Pelle

FBgn0010441

cn1855

Tube

FBgn0003882

Gmsg-10360

MyD88

FBgn0033402

cn12871

PGRP-LC (peptidoglycan recognition protein LC) PGRP-LE (peptidoglycan recognition protein LE) PGRP-LF (peptidoglycan recognition protein LF) PGRP-SA (peptidoglycan recognition protein SA) PGRP-SD (peptidoglycan recognition protein SD) Imd (immune deficiency) dTAK1 (TGF-beta activated kinase 1) Kenny (IKKg) (IkappaB kinase) Dredd (Death released ced-3/NeddZ-like protein) Dnr1 (defence repressor 1)

FlyBase ID

cn16500 cn353

Gmsg-10321 cn14933

FlyBase IDa

Orthologb

FBgn0035976

cn1360

FBgn0030695

Gmm-3156

FBgn0035977

cn1360

FBgn0030310

Tse122g03

FBgn0035806

cn1360

FBgn0013983 FBgn0026323

cn3932 Gmsg-7119

FBgn0041205

Gmsg-7350

FBgn0020381

cn13356

FBgn0260866

Tse104g08.qlc (continues)

TABLE 1

(Continued)

Toll Drosophila Molecule

FlyBase IDa

Imd Orthologb

Cactus

FBgn0000250

Gmsg-2191

Dorsal

FBgn0260632

cn12921

Dif (dorsal-related immunity factor) DaPKC (Drosophila atypical protein kinase C) WntD (Wnt inhibitor of Dorsal)

FBgn0011274

Gmsg-2191

FBgn0022131

cn10314

FBgn0038134

Not identified

a b

Drosophila Molecule DIAP2 (inhibitor of apoptosis-2) Ird5 (IKKb) (Immune response deficient 5) dFADD

FlyBase IDa

Orthologb

FBgn0015247

cn3040

FBgn0024222

cn5893

FBgn0038928

cn6552

dTAB2 (TAK1-associated binding protein 2)

FBgn0086358

Gmsg-6058

Caspar

FBgn0034068

GLAEJ30TV

Relish Caudal

FBgn0014018 FBgn0000251

cn7095 cn9066

Drosophila melanogaster accession numbers from genome data assembled by Flybase http://flybase.org. Glossina orthologs obtained from G. m. morsitans EST libraries clustered by GeneDB http://www.genedb.org/gendb/glossina.

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from a microbicidal to a recognition role (Lemaitre and Hoffmann, 2007). A role for PGRP-LB, a known negative regulator of the Imd pathway in Drosophila, has been noted in downregulating tsetse immune responses to virulent but not avirulent bacteria (Weiss et al., 2008). Notably, no PRR has yet been identified for African trypanosomes, although it is clear that there is a tsetse immune response to their presence (Hao et al., 2001). Binding of PRRs to a pathogen leads to activation of Toll/Imd downstream signalling cascades. Signalling culminates in the activation and nuclear translocation of NFkB transcription factors Dorsal/Dif (Toll pathway) or Relish (Imd pathway). This in turn causes transcription of immune effector genes such as HDPs. Importantly, molecular crosstalk may occur between the Toll and Imd pathways, thus giving rise to a wide spectrum of possible immune responses to pathogen invasion (Tanji and Ip, 2005; Tanji et al., 2007). In Drosophila, control of induced epithelial immune responses is associated with the Imd signalling pathway (Tzou et al., 2000). Gene knockdown of the Imd pathway transcription factor Relish in tsetse flies has been demonstrated to cause an increase in midgut and salivary gland infection rates with trypanosomes (Hu and Aksoy, 2006). This evidence implicates the Imd pathway in regulating fly susceptibility to trypanosome infection. Furthermore, knockdown of Relish leads to downregulation of attacin, a HDP associated with tsetse– trypanosome interactions (Hao et al., 2001; Hu and Aksoy, 2005, 2006; Nayduch and Aksoy, 2007).

2.5.4

Effector molecules

HDPs are evolutionarily conserved effector molecules of the humoral defence system and are found among all classes of life. They exhibit a broad spectrum of activity against bacteria, fungi, viruses and transformed cells (Lemaitre and Hoffmann, 2007). In addition, the anti-parasite activity of HDPs has been illustrated in several vector–parasite systems (Durvasula et al., 1997; Shahabuddin et al., 1998; Boulanger et al., 2002b) including a tsetse– trypanosome system (Hu and Aksoy, 2005; Hu and Aksoy, 2006). Many HDPs target pathogens by disturbing the pathogen membrane potential or by disrupting internal cell functioning leading to cell death by apoptosis or necrosis. Early research into these immune mediators in G. m. morsitans identified four HDPs: an attacin (AttA1), a cecropin, a defensin and a diptericin (Hao et al., 2001; Boulanger et al., 2002a). More recently, characterization of the G. m. morsitans attacin loci has recognized that attacin genes are organized in three clusters encoding three different attacins: attA, attB and attD. The amino acid sequences of AttA and AttB are almost identical while AttD is only 69% identical to the AttA/B form. These genes are differentially regulated (Wang et al., 2008). Interestingly, two additional HDPs, one with anti-Gram-negative activity and the other with anti-Gram-positive activity, have also identified

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following trypanosome challenge (Boulanger et al., 2002a). These HDPs remain uncharacterized. In vivo analysis of HDP transcript expression during trypanosome infection has indicated that these peptides are differentially regulated in the haemolymph and in the major immunoresponsive tissues of midgut, fat body and proventiculus (Hao et al., 2001, 2003; Hu and Aksoy, 2006). Early in the infection process, the presence of trypanosomes in the midgut or haemolymph does not lead to activation and increased transcription of midgut or fat body HDP genes (Hao et al., 2001). However, by day 6, as parasite numbers increase, attacin (AttA/B and AttD) and defensin transcript expression is high in the fat body (Hao et al., 2001, 2003; Wang et al., 2008). In selfcured flies HDP transcript expression levels fall, but in flies with established midgut infections expression levels remain high in the fat body and proventriculus (Hao et al., 2001, 2003). This does not appear to affect the viability of the parasite population within the midgut, although it remains to be seen if individual parasites are affected, thus resulting in a change in the nature of the parasite population. It is possible that trypanosomes exhibit a stage-specific sensitivity to particular immune molecules, with procyclics exhibiting higher resistance to the trypanocidal activity of HDPs than BSF trypanosomes as was observed in vitro by Haines et al. (2003). Alternatively fat body synthesized peptides circulating in the fly haemolymph may fail to reach parasites located in the midgut environment. The trypanocidal activity of HDPs has been recognized. Stomoxyn, isolated from the facultative hematophagous fly Stomoxys calcitrans, exhibits trypanocidal activity (Boulanger et al., 2002b). In addition, there is direct evidence of trypanosome killing by Glossina HDPs themselves (Hu and Aksoy, 2005; Hu and Aksoy, 2006; Nayduch and Aksoy, 2007). Recombinant attacin (AttA1) inhibited both BSF and PCF growth in vitro (Hu and Aksoy, 2005). In vivo, gene knockdown of the AttA1 peptide, or its transcriptional regulator Relish, led to a statistically significant increase in midgut and mature salivary gland trypanosome infection rates (Hu and Aksoy, 2006). Relish also regulates cecropin expression in Glossina, but whether cecropin possesses trypanocidal properties is yet to be directly investigated. More recently, differential expression of AttA1 transcripts in a trypanosome susceptible species (G. m. morsitans) and two comparatively trypanosome-refractory species (G. pallidipes and G. p. palpalis) has been reported (Nayduch and Aksoy, 2007). Refractory species showed higher attacin transcript expression in fat body (systemic) and proventiculus/ midgut (local) tissues in comparison to susceptible flies in both teneral and blood-fed states. Knockdown of attacin expression in G. pallidipes led to increased trypanosome susceptibility, although the possible confounding effects of high mortality rate, starvation, wounding and low sample number in this study should be noted (Nayduch and Aksoy, 2007). Nevertheless, the evidence suggests that attacin may be an important regulator of tsetse–trypanosome interactions.

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Notably, several investigations of HDP function in tsetse–trypanosome interactions bypass a natural trypanosome developmental stage by using PCF rather than BSF trypanosomes to infect the flies (Hao et al., 2001, 2003). It is too early to say how this experimental approach to infection affects the tsetse– trypanosome interaction but it clearly runs the risk of bypassing part of the natural fly immune response. In some cases, the authors do not state the trypanosome life cycle stage used (Nayduch and Aksoy, 2007). Furthermore, supplementation of bloodmeals with glucosamine (Hao et al., 2001, 2003) may disrupt the natural infection process and confound the interpretation of results (Peacock et al., 2006). The HDP diptericin is constitutively expressed in the proventriculus and fat body of tsetse (Hao et al., 2001). This HDP expression profile has been attributed to the presence of symbionts, in particular to the presence of Sodalis in the gut and haemolymph. Interestingly, Sodalis is resistant to the Gramnegative bactericidal activity of mature synthetic diptericin in vitro (Hao et al., 2001), whereas the non-native bacterium E. coli is not. Sodalis is also more resistant to recombinant attacin (recGmAttA1) than E. coli (Hu and Aksoy, 2005) and to a battery of other Gram-negative and Gram-positive bacteriolytic HDPs (Haines et al., 2003). Thus, it is possible Sodalis may have evolved HDP resistance traits permitting survival in the hostile tsetse midgut environment while invading pathogens are eliminated. 2.5.5

Antioxidants

In addition to the NFkB pathway-mediated defense systems, the NFkB-independent production of microbicidal ROS is a key component of insect epithelial immune responses (Ha et al., 2005a; Lemaitre and Hoffmann, 2007). In Drosophila, natural gut infections with bacteria are associated with rapid synthesis of ROS. Studies have demonstrated that it is Drosophila dual oxidase (dDuox), and not NADPH oxidase (dNox), that provides the main source of ROS that limits bacterial proliferation in the midgut (Ha et al., 2005a,b). The infection-inducible nature of intestinal dDuox suggests the transcriptional role of dDuox may play a pivotal role in fly midgut protection against invading pathogens. dDuox is capable of generating H2O2, and the highly microbicidal HOCl is derived from H2O2 by neutrophil-derived myeloperoxidase (MPO) type activity. Physiological regulation of ROS levels in the midgut is achieved via dDuox-dependant ROS generation and immune-regulated catalase (IRC)dependant ROS removal (Ha et al., 2005a,b). A fine redox balance is critical, as knockdown of either dDuox or IRC leads to higher mortality rates due to either insufficient or prolonged oxidative stress responses respectively. It is known that ROS activates a cell death pathway in procyclic T. b. brucei trypanosomes (Ridgley et al., 1999). In tsetse, increased ROS and nitric oxide (NO) transcript expression have been reported in the proventriculus in response to trypanosome challenge (Hao et al., 2003). Additionally, upregulation of

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oxidative stress genes has been demonstrated in the midgut transcriptome of infected flies (Lehane et al., 2003; Munks et al., 2005) and self-cured flies (Lehane et al., 2008). Hao et al. have hypothesized that reactive intermediates such as NO or H2O2 may function as chemical signals to mediate communication between different immunoresponsive tissues (Hao et al., 2001, 2003). Indirect evidence suggests a role for antioxidants in signalling during parasite development in the fly and protecting tsetse flies against trypanosomes (Hao et al., 2001, 2003; Macleod et al., 2007a,b). Recently, Macleod et al. (2007b) demonstrated the feeding of different antioxidants (glutathione, cysteine, N-acetyl-cysteine, ascorbic acid or uric acid) in the infective bloodmeal led to significant increases in midgut infection rates. Also, glucosamine, which has routinely been used to boost trypanosome midgut infection rates, has been shown to be an antioxidant molecule (Xin et al., 2006). The original interpretation of increased infection rates in flies fed glucosamine was that this sugar neutralized specific trypanocidal lectins in the midgut, permitting higher infection rates to occur (Maudlin and Welburn, 1987; Welburn et al., 1989; Murphy and Welburn, 1997) (see Section 2.5.6). However, with this more recent demonstration that N-acetyl glucosamine can scavenge ROS, superoxide and hydroxyl ions (Xin et al., 2006), an effect on trypanosome infection rates through reduction of ROS challenge is a strong alternative hypothesis. Oxidative stress is believed to be involved in control of parasites in other vector–parasite systems, including the Plasmodium–mosquito and Trypanosoma–Rhodnius systems. Augmented production of NO has been noted in Anopheles stephensi, An. gambiae and An. pseudopunctinpennis following P. berghei infection (Luckhart et al., 1998; Herrera-Ortiz et al., 2004). The R. prolixus NO system responds to T. rangeli and T. cruzi, implicating involvement in trypanosome infection regulation (Whitten et al., 2001, 2007). Furthermore, the stable suppression of trypanosome parasitaemia in Cape buffalo is associated with increased levels of serum ROS (Wang et al., 2002). A role for transferrin in immune signalling and the upregulation of NO in vertebrates has been suggested (Stafford and Belosevic, 2003). Recently, knockdown of transferrin in G. m. morsitans was demonstrated to have a statistically significant impact on trypanosome prevalence, resulting in almost a doubling of trypanosome midgut infection rate (Lehane et al., 2008). The mechanism of involvement of tsetse transferrin in mediating tsetse– trypanosome interactions is still unknown, but there is the intriguing possibility that it may be analogous to the function in vertebrates. 2.5.6

Lectins and programmed cell death

Susceptibility, defined as midgut establishment of T. congolense or T. b. brucei in G. m. morsitans infected at the first bloodmeal, can be selected for with results apparent in the F1 generation (Maudlin, 1982). This susceptibility is an extra-chromosomal trait inherited through the female line (Maudlin and

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Dukes, 1985). G. m. centralis infected with T. congolense show similar susceptibility phenotypes. It appears the effect operates in both the midgut and foregut, because selected lines showed similar phenotypes to T. vivax infection (Moloo and Kutuza, 1988a). The lectin hypothesis was proposed as the mechanism underpinning this phenotype by Maudlin and Welburn (1987). They argued that midgut-expressed lectins were the predominant factor killing trypanosomes in vivo in the tsetse fly, with lectin levels modulated by the changing number of the symbiont Sodalis glossinidius in the fly midgut (Welburn and Maudlin, 1999). This argument was based primarily on evidence that feeding of sugars capable of inhibiting lectins increased trypanosome midgut infection rates (Maudlin and Welburn, 1987; Ingram and Molyneux, 1988; Welburn et al., 1994). Additionally, based on the effects of the plant lectin concanavalin A (Con A) on trypanosomes in vitro, lectin-mediated killing of trypanosomes was believed to occur by a process termed proto-apoptosis (Welburn and Maudlin, 1999; Pearson et al., 2000) (see below). A lectin specific for D-glucosamine and with lesser affinity for N-acetyl-D-glucosamine has been tentatively identified in tsetse midgut (Ibrahim et al., 1984; Ingram and Molyneux, 1988). Most of this lectin is found attached to the PM (Lehane and Msangi, 1991). Inhibition of midgut lectins was hypothesized to occur naturally to the greatest extent in newly emerged flies, where lectins were neutralized by the build-up of inhibitory sugars released from chitin by the endochitinase of the symbiont Sodalis during the pupal period (Welburn and Maudlin, 1999). The increased potential of older starved flies to be infected when compared with age-matched non-starved flies (Kubi et al., 2006) is in disagreement with the lectin hypothesis, as both are reported to have similar midgut lectin levels (Lehane and Msangi, 1991). More recently, the multiple effects of N-acetyl-Dglucosamine or D-glucosamine inclusion in the bloodmeal upon trypanosome growth and tsetse physiology have been reported (Peacock et al., 2006; Ebikeme et al., 2008). Both sugars slowed bloodmeal movement along the midgut. Glucosamine significantly increased the size of bloodmeal taken, as well as increasing the fly mortality rate. Interestingly, N-acetyl-D-glucosamine stimulated trypanosome growth in the midgut and in vitro in the absence of any fly-derived factors. The ability of N-acetyl-D-glucosamine to enhance trypanosome PCF growth in vitro does not seem to involve any direct effect of N-acetyl-D-glucosamine upon metabolic processes within the trypanosome (Ebikeme et al., 2008). In vitro experiments have instead shown that N-acetylD-glucosamine indirectly enhances the rate of L-proline metabolism by inhibiting the trypanosome hexose transporter, thus depriving the parasite of D-glucose (Ebikeme et al., 2008). However, PCFs cultured in D-glucose free medium, and thus possessing a metabolism primed for L-proline use, do not appear to have a higher infection rate when used to infect tsetse flies. This suggests that the metabolic impact of N-acetyl-D-glucosamine on trypanosomes has no bearing on the success of trypanosome establishment within the tsetse midgut (Ebikeme et al., 2008). These off-target effects of feeding glucosamine complicate the

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interpretation of experiments on the trypanosome prevalence phenotype observed (Peacock et al., 2006). Above all, the demonstration that glucosamine (which has routinely been included in infected bloodmeals to artificially boost trypanosome midgut infection rates) is an antioxidant molecule (Xin et al., 2006) suggests a simpler explanation (see Section 2.5.5) for the effect of this sugar on tsetse–trypanosome interactions. In addition, the lectin hypothesis contends that higher symbiont densities lead to increased susceptibility. However, a correlation between symbiont density and susceptibility does not occur in all instances (Shaw and Moloo, 1991; Weiss et al., 2006). Successfully established midgut infections show an average parasite density of approximately 5  105 cells per midgut (Van den Abbeele et al., 1999; Gibson and Bailey, 2003), but this number can vary with clone and species of trypanosome. Procyclic trypanosomes share a common energy source (proline) with their tsetse hosts (Welburn and Maudlin, 1999), which may become a limiting factor if trypanosome numbers expand too far. It is possible that trypanosomes self-regulate their numbers and that quorum sensing may be involved in the process (Acosta-Serrano et al., 2001; Roditi and Lehane, 2008). More specifically, it has been suggested that the cleaved N-terminal fragments of trypanosome procyclins might be involved in a density-sensing mechanism, which the trypanosomes use to control their population densities (Acosta-Serrano et al., 2001). However, this mechanism may not be essential as procyclin knockout mutants are still fly-transmissible (Vassella et al., 2009). A novel form of cell death is believed to occur in trypanosomes, known as proto-apoptosis, and it is suggested that this is part of the mechanism by which trypanosomes regulate numbers within the midgut (Welburn et al., 1996; Murphy and Welburn, 1997; Welburn and Maudlin, 1997). In support of this idea, incubation of PCFs of T. b. brucei, T. b. rhodesiense or T. congolense trypanosomes (but not their corresponding BSFs) with the plant lectin Con A in vitro induces growth arrest and death, with many of the characteristics of programmed cell death (PCD) (Welburn and Maudlin, 1997; Murphy and Welburn, 1997). In tsetse, with some trypanosome infections, cyst-like forms have been observed embedded in the PM (Robertson, 1913; Evans and Ellis, 1983; Gibson and Bailey, 2003). Some cysts contain highly motile forms while others appear to consist of aggregates of rounded up trypanosomes. Whether these cysts represent an unknown life cycle stage or degenerating trypanosome forms is unknown. Gibson and Bailey (2003) noted that these forms resembled the morphologically altered PCF trypanosomes treated with Con A in vitro to induce cell death. Proteins upregulated during the process of PCD include the trypanosome protein kinase C receptor, which is similarly upregulated in the short stumpy BSF. This suggests that stumpy form BSFs are removed from the trypanosome population within the mammalian hosts (should ingestion by a tsetse host not occur). This would avoid triggering a wider immune response within the

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mammalian host, which could jeopardize the survival of the long slender BSF population (Murphy and Welburn, 1997). 2.5.7

Other potential effector molecules

Melanization plays an important role in insect defence reactions such as wound healing, sequestration of microorganisms, encapsulation and the production of intermediates toxic to invading pathogens (Lemaitre and Hoffmann, 2007). Prophenoloxidases (ProPOs) are oxidoreductases related to hemocyanins that mediate melanization. These molecules have been implicated in tsetse– trypanosome interactions as inhibition of phenoloxidase (PO) activity with phenylthiourea (PTU) increases midgut trypanosome infection rates (Nigam et al., 1997). Furthermore, T. b. rhodesiense has been shown to significantly activate haemolymph proPO of female G. m. morsitans (Nigam et al., 1997). Such a complex biochemical cascade as occurs in the ProPO system is almost certainly highly reliant on the controlled environment of the haemolymph for its successful operation, and it is hard to see how it could operate effectively in the midgut lumen, with the possible exception of the immediate surface of the epithelium. However, it is possible that proPO may be a means of killing those parasites which do enter the haemocoel of the fly. TsetseEP protein may also play a role in tsetse–trypanosome interactions. This immunoresponsive protein is expressed strongly in the midgut of Glossina (Chandra et al., 2004) and is upregulated in response to immune stimulation with Gram-negative bacteria (Haines et al., 2005). This protein exhibits a high sequence identity to the carboxy terminal region of the EP form of procyclins, which is found on the surface of procyclic T. b. brucei. The glutamic acid– proline (EP) repeats found on both molecules are extremely unusual and it seems unlikely that they coincide in the tsetse midgut solely by chance. The potential role of tsetseEP in mediating fly responses to trypanosome infection merits further investigation. 2.5.8

Adaptive immunity

The existence of immunological memory has long been known in vertebrates, with humans utilizing vaccination before key mechanisms of adaptive immunity were known or understood. Vertebrate adaptive immunity is characterized by long-term protection against specific antigens achieved in part by antibodies that consist of a large diversity of somatically rearranged immunoglobulin receptors. The possibility that invertebrates are capable of adaptive immunity to pathogen challenge has recently re-emerged as a possibility (Watson et al., 2005; Dong et al., 2006; Dong and Dimopoulos, 2009). For example, Dong et al. (2006) identified Down syndrome cell adhesion molecule (Dscam) as an alternatively spliced, hypervariable immunoglobulin domain containing gene responsible for generating a broad range of pathogen recognition receptors

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(PRRs) in Anopheles gambiae mosquitoes. The gene contains 101 exons including three variable Ig exon cassettes. Alternative splicing of this gene produces a highly diverse set of over 31,000 potential splice forms. Challenge with different pathogens leads to production of specific AgDscam splice-form repertoires. Interestingly, an ortholog of Dscam has been identified from a Glossina EST library (cn9121, e-value 6.0  10 21). Whether Glossina uses this gene in immune responses to trypanosomes or bacteria remains to be elucidated. 2.5.9

Fly immune system and microbial balance

Glossina carries a range of microorganisms (see Section 3) and hence, must manage to coexist with these symbionts while still protecting themselves from the onslaught of pathogens. The host factors that maintain the homeostatic relationship between the tsetse host and its symbionts are largely unknown, and our current insight is based primarily on studies in Drosophila. Recently, the intestinal homeobox gene Caudal (Cad) has been identified as a key factor in repressing NFkB-dependant HDP genes in Drosophila (Ryu et al., 2008). Thus, although Drosophila commensal organisms can induce a high level of local Imd-regulated NFkB activation, only a subset of target genes is activated. Regulation of the immune system is tightly controlled, as overexpression of HDP genes in Caudal knockdown flies caused a shift in the commensal population in the midgut. In particular, the dominance of Gluconobacter sp. strain EW707 eventually led to gut apoptosis and host mortality in Drosophila. The model proposed by Ryu et al. (2008) involves interplay between Caudal and the NFkB transcription factor Relish to regulate the expression of HDPs, which in turn defines the microbial community and insect health. Whether this is also the case in Glossina has not yet been investigated, although an ortholog of Caudal has been identified in the EST libraries (cn9066, e-value 1.2  10 47) making this a possibility. 2.6

EFFECTS OF TRYPANOSOME INFECTION ON TSETSE PHYSIOLOGY

Trypanosome infection may have multiple effects on the tsetse host. A suppression subtractive hybridization study (Lehane et al., 2008) identified molecules differentially expressed in established versus self-cured T. b. brucei infections in G. m. morsitans. Analysis of the gene fragments generated suggested that trypanosome infection has a marked effect upon the metabolism of the fly host. Flies self-cured of trypanosome infection displaying the signals of increased energy usage and an oxidative stress response compared to infected flies. Within the tsetse fly, procyclic trypanosomes employ the amino acid proline as a source of energy, while the fly typically uses proline reserves for flight. In infected male flies, trypanosomes infection can reduce the flight potential by 15% (Bursell, 1981). Trypanosome infection in female flies would be far more

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costly in terms of flight potential as females have smaller proline reserves due to the energy required for larval development. Recently, Hu et al. (2008) investigated the cost of trypanosome infection on fly reproductive fitness. Two trypanosome strains were selected that differentially activated the host immune system: a T. b. rhodesiense wild-type strain that induced expression of attacin and defensin in trypanosome infected flies and a mutant strain that did not. Only infection with the wild-type strain led to a significant increase in larval deposition periods and a decrease in milk gland protein expression. This suggests that activation of tsetse immune responses by infection with immunogenic trypanosomes delays larvigenesis via decreased expression of the milk gland protein vital for larval growth. Trypanosome infection may also impact the feeding behaviour of infected tsetse hosts. Infection of G. m. morsitans and G. austeni with T. b. brucei reportedly results in increased probing behaviour and more voracious feeding (Jenni et al., 1980). Accumulation of large numbers of trypanosomes in the salivary glands or proboscis has been observed, and may impede function of labral mechanoreceptors as detectors of blood flow rate in the gut (Molyneux and Jenni, 1981). Thus an infected fly may take several smaller meals or feed for longer, either scenario potentially leading to increased trypanosome transmission to the vertebrate host.

2.7

FLY SEX, AGE AND STARVATION AND TRYPANOSOME TRANSMISSION

Susceptibility to parasite infection is also influenced by the sex of the fly. This is evident in midgut infections with T. congolense and T. brucei, but not in infections with T. vivax. Most laboratory studies (Burtt, 1946b; Harley, 1971; Distelmans et al., 1982; Mwangelwa et al., 1987; Hide et al., 1991; Moloo, 1993; Welburn et al., 1995; Dale et al., 1995), but not all (Welburn and Maudlin, 1992; Moloo et al., 1992; Moloo, 1993), suggest that male flies are more susceptible. Female G. p. palpalis are more resistant to developing mature T. congolense infections (Distelmans et al., 1982) than males. Unexpectedly, this sex-dependent phenomenon was not observed with T. congolense infections in G. m. morsitans (Dale et al., 1995). Of interest, compared to a virgin G. m. morsitans, a mated female is twice as refractory to T. b. brucei infection (Macleod et al., 2007a). Although both sexes are capable of transmitting parasites, both male and female flies are innately resistant to parasite infection and this resistance increases rapidly with age (Fig. 4). This natural refractoriness to trypanosome maturation and/or establishment is seen in both laboratory and field populations (Distelmans et al., 1982; Leak 1999) and is termed the teneral phenomenon (Welburn and Maudlin, 1992). In the tsetse community the term teneral is used to describe a newly emerged fly that has not yet had its first bloodmeal. The fly during this time is marked by

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a soft, often described as soapy-feeling, exoskeleton and lighter body coloration. While tsetse researchers universally use the word ‘‘teneral’’, they seem to have forgotten that the term represents a highly variable physiological state, particularly in terms of fly susceptibility. Depending on tsetse species and the nutritional status of the newly eclosed adult fly, teneral flies can survive for up to 1 week before dying of starvation (Kubi et al., 2006; Lehane lab, unpublished observations). Van Hoof et al. (1937) originally demonstrated the ‘‘teneral phenomenon’’, that is the higher susceptibility of a newly emerged fly to trypanosome infection compared to an older fed fly, using G. f. fuscipes challenged with T. b. gambiense. The highest midgut establishment rates were obtained in flies fed the infective meal 0–1 days post-emergence (p.e.). This work was later confirmed by Wijers (1958), who observed that G. p. palpalis emergents less than 30 h p.e. were more susceptible to a first infective bloodmeal infection than flies 30–78 h p.e. In recent years, the teneral definition has shifted to a nutritional focus and teneral has become synonymous with ‘‘unfed’’. Many published papers ignore the fact that parasite susceptibility during this teneral period can vary widely, and consequently flaws in experimental design may confound data interpretation. Speculation on why newly emerged flies are so susceptible to trypanosomes is profuse. Incomplete formation of the PM, variable antioxidant or lectin concentrations, molecular maturity of the midgut (i.e. proteases), immaturity of the immune response, larval meal turnover (preand post-meconium expulsion), pH gradient differences and variation in symbiont loads may, either singly or as a combination, play a role in creating the teneral phenomenon. Although further research is required to confirm which of these factors (if any) contribute to this age-related susceptibility to infection, it is crucial to standardize trypanosome infection experiments by redefining the definition for teneral as ‘‘hours post-eclosion’’ instead of simply ‘‘unfed’’. Starvation is a constant risk the tsetse fly faces throughout its lifetime. Flies deprived of a bloodmeal utilize energy reserves stored in the fat body and risk death when fat reserves drop to approximately 6% of the total dry body-mass (Rogers et al., 1994). Male flies, which use most of each bloodmeal for lipid production, are more resistant to starvation than females, which use the bloodmeal for both lipid production and growth of the larva (Randolph and Rogers, 1981). Starvation may influence feeding behaviour with the range of host choice expanding as the fly’s hunger intensifies (Bouyer et al., 2007) with obvious consequences for the epidemiology of disease. Environmental conditions, such as temperature and relative humidity, also influence fly starvation with many Glossina species becoming hungrier during the dry season (Jackson, 1933). Starvation is a key factor in tsetse–trypanosome interactions because the nutritional status of the fly at the time of the infective bloodmeal is a key determinant of fly susceptibility to trypanosome infection (Gingrich et al., 1982; Mwangelwa et al., 1987; Gooding, 1988; Kubi et al., 2006).

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Under laboratory conditions with bloodmeals given every 4 days, newly emerged male tsetse flies are highly susceptible to trypanosome infection at the first bloodmeal while older flies exhibit a significantly lower susceptibility to trypanosome infection (Fig. 4) (Distelmans et al., 1982; Welburn and Maudlin, 1992; Kubi et al., 2006). However, field studies of trypanosome prevalence with fly age suggest a significantly higher proportion of the older adult tsetse population than predicted develop a mature trypanosome infection (Woolhouse et al., 1993; Msangi et al., 1998; Lehane et al., 2000). Recently, laboratory-based studies by Kubi et al. (2006) demonstrated a period of starvation (3–4 days for teneral or 7 days for adult flies) increased fly susceptibility to T. b. brucei or T. congolense infection, resulting in a significantly higher number of T. congolense or T. b. brucei mature salivary gland infections. In the case of T. congolense, a significantly higher number of midgut infections were also observed. It is likely that the increased infection rates noted in older field flies compared to laboratory flies is explained by starvation events in the field. An explanation for this observation may be that under high nutritional stress the physiological barriers to trypanosome infection are suppressed because the energy cost of an immune response is high (Schmid-Hempel, 2005; Ye et al., 2009). Trypanosome infection may have a negative impact on fly longevity (Bursell, 1981). More immunogenic lines of trypanosomes adversely affect tsetse reproductive fitness (Geiger et al., 2008), which suggests that trypanosome infection places a selection pressure on tsetse flies.

2.8

ENVIRONMENTAL TEMPERATURE AND TRYPANOSOME TRANSMISSION

As puparial incubation temperature rises (within biological limits), so does the ability of the corresponding adult fly to develop a mature infection (Burtt, 1946a; Ndegwa et al., 1992). This is not attributed to an increase in the number of flies that establish a midgut infection, but rather to an increase in the proportion of these flies that go on to develop mature infections (Dipeolu and Adam, 1974). The mechanism responsible is unknown. However, this observed trend may have consequences in the field for the distribution of trypanosomiasis. There is a positive correlation between latitude relative to 7  S (the median of tsetse distribution) and infection rate (Ford and Leggate, 1961). Although the factors involved in the field are undoubtedly complex (Jordan, 1965), ambient temperature also increases towards the 7  S median, implying that it may be puparial incubation temperature that influences this trend. If that is the case, then temperature plays a central role in determining the geographical distribution and epidemiology of trypanosomiasis.

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Symbiont–tsetse–trypanosome interactions WIGGLESWORTHIA GLOSSINIDIUS

All Glossina sp. harbour W. glossinidius, a primary obligate (beneficial) Gramnegative endosymbiont. Based on concordant evolutionary analyses, it is estimated the Wigglesworthia-Glossina symbiosis evolved in ancestral forms between 50 and 80 million years ago (Chen et al., 1999). This bacterium resides within the cytoplasm of specialized epithelial midgut cells, bacteriocytes, which form a horseshoe shaped organ, the bacteriome (Fig. 2E), located in the anterior midgut (Aksoy et al., 1995; Aksoy, 1995). The Wigglesworthia genome is grossly reduced in size to less than 700 kb and is predicted to encode for 617 proteins (Akman et al., 2002). This streamlined genome has undergone massive gene erosion as genes required for defense mechanisms, metabolic pathways, DNA repair mechanisms and so on, are no longer essential as this endosymbiont exists in a stable host environment. In fact, even a gene involved in chromosome replication, dnaA, which is considered essential for bacterial survival, has been lost in Wigglesworthia (Akman et al., 2002). This member of the Enterobacteriacae family, first observed over a century ago (Stuhlmann, 1907), was thought to produce metabolites to compensate for nutritional deficits in the host’s haematophagous diet and appears to be partially associated with the metabolism of B-complex vitamins essential for tsetse survival (Wigglesworth, 1929; Nogge, 1981; Akman et al., 2002). The elimination of Wigglesworthia reduces fly longevity, rate of bloodmeal digestion and fecundity by making females sterile (Nogge, 1976; Dale and Welburn, 2001; Pais et al., 2008), although supplementation of the host bloodmeal with vitamins partially reverses this infertility phenotype (Nogge, 1981). It has been suggested that the bacterium may also influence host digestive processes through either synthesis of protoheme or glycerophospholipids, fatty acids and steroids (Pais et al., 2008). Wigglesworthia is vertically transmitted from the female fly to her progeny via milk gland secretions (Denlinger and Ma, 1975). More specifically, in situ hybridization has identified Wigglesworthia exclusively associated with the milk gland lumen and the canals leading to the secretory reservoirs (Attardo et al., 2008). Recently, studies by Pais et al. (2008) have demonstrated that selective elimination of Wigglesworthia symbionts by ampicillin antibiotic treatment leads to increased susceptibility to trypanosome midgut infection in non-teneral flies. This implies that symbiont clearance may lead to a decrease in fly basal immunity, thus affecting host immune responses to trypanosome infection. However, these Wiggleworthia-cleared flies also had a compromised ability to digest their bloodmeals and increased trypanosome susceptibility was only observed in older flies. Thus, neither the effect of ‘‘starvation’’, known to cause increased parasite susceptibility (Kubi et al., 2006), nor the effects of antibiotic treatment on the fly and its other symbionts can be discounted.

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WOLBACHIA PIPIENTIS

A secondary tsetse symbiont, Wolbachia pipientis, is also maternally inherited. Unlike Wigglesworthia, it colonizes the female reproductive organs (not the milk gland) and is transovarially transmitted to progeny. Wolbachia are the most frequently described symbiotic association in arthropods; surveys have indicated that more than 70% of all insect species are infected with Wolbachia (Werren and Windsor, 2000). Wolbachia belong to the subdivision of Gramnegative intracellular alpha-proteobacteria and are most famous for their reproductive manipulations of the host including parthenogenesis, feminization, male-killing and cytoplasmic incompatibility (for a review of Wolbachia insect symbioses see Siozios et al., 2008). It has been reported that 100% of laboratory-reared tsetse colonies are infected with Wolbachia, while infections in wild populations vary significantly (Cheng et al., 2000). Within infected tsetse, Wolbachia colonizes several tissues, depending on the species of Glossina. Using a Wolbachia-specific PCR-based assay, Wolbachia was found in the reproductive tissues of all infected tsetse species examined. However, a wider tissue tropism was observed in the somatic tissues (gut, head, muscle, fat body, milk gland and salivary gland) in certain populations of G. austeni (Cheng et al., 2000). The role Wolbachia plays in the tsetse is unknown and whether its presence influences trypanosome infection rates by stimulating the fly immune system remains unknown. There are strong incentives to investigate whether or not Wolbachia induce cytoplasmic incompatibility in tsetse because it is one means by which selective traits can be driven through insect populations. As such, it is a highly desirable tool for promoting the success of paratransgenesis, a potential disease control strategy involving engineered symbionts and alteration of fly susceptibility to parasite infection – see Section 4.2 (Aksoy et al., 2003; Aksoy and Rio, 2005). 3.3

SODALIS GLOSSINIDIUS

The third tsetse symbiont, S. glossinidius, is a Gram-negative, microaerophilic, non-motile bacterium that forms a new bacterial taxon and species in the family Enterobacteriaceae (Dale and Maudlin 1999). Reinhardt et al. (1972) first described Sodalis as small rickettsia-like organisms (RLO) that were separated from the cytoplasm of midgut cells by a clear lytic zone. Pinnock and Hess (1974) confirmed these observations and reported the presence of this pleomorphic microbe in other tissues such as the fat body and ovaries. The development of a Sodalis-specific PCR-assay further established that this symbiont resided in the midgut, haemolymph and milk gland of teneral flies (Cheng and Aksoy, 1999; Attardo et al., 2008). Sodalis can live both intracellularly and as free-living forms in the gut lumen. This bacterium is one of the only insect symbionts that has been adapted to

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in vitro culture (Dale and Maudlin, 1999; Matthew et al., 2005), making it a model organism particularly for research on host cell invasion, evolutionary biology (the transition from free-living to obligate endosymbiosis) and phylogenetic associations. In recent years, two new symbionts belonging to the Sodalis lineage have been identified, a bacterial isolate from the bloodsucking fly, Craterina melbae (Novakova and Hypsa, 2007) and a bacteriocyteassociated symbiont from the chewing louse, Columbicola columbae (Fukatsu et al., 2007). Unlike other bacterial symbionts, Sodalis does not produce catalase (characteristic of several microaerophiles) which, when combined with the results of additional phenotypic tests, indicates that Sodalis has a reduced biochemical profile when compared to other members of the family Enterobacteriaceae (Dale and Maudlin, 1999). Initially, 85% of the Sodalis genome was characterized by hybridization to E. coli gene macroarrays (Akman and Aksoy, 2001) and many potential gene products were identified. The use of E. coli arrays of course did not allow the identification of molecules unique to Sodalis, as exemplified by the failure to detect genes of the type III secretion system (Dale and Welburn, 2001) and genes encoding two chitinases. In 2006, the genome of Sodalis was published (Toh et al., 2006). Unlike Wigglesworthia, Sodalis has a much larger genome (4.1 Mb), but surprisingly it is predicted to encode only 2432 proteins (which is approximately 50% of what would be expected of a free-living organism). The presence of 972 pseudogenes indicates this genome is gradually eroding, although a previously designated pseudogene (carA) has now been shown to be functional and transcriptionally responsive despite its truncation (Pontes et al., 2008). Extrachromosomal DNA revealed another 160 potential ORFs located on four circular elements (Darby et al., 2005). While this would indicate a general transition to an obligate mutualistic association with the tsetse, several virulence related-factors such as haemolysin, phospholipases and invasion proteins have maintained functionality, implying that pathogenic genes are being retained (Dale and Welburn, 2001; Toh et al., 2006; Feldhaar and Gross 2009). Observations by Pinnock and Hess (1974) support this virulence concept as originally Sodalis was described as a pathogen based on electron microscopy of infected tsetse tissues; the host lytic zones surrounding Sodalis might be indicative of an adverse reaction between the tsetse host and the bacteria. In addition, the highly infected fat body in Glossina pallipides showed signs of cellular disruption and degeneration that could not be explained as procedural artefacts. This cellular disruption was also observed when Sodalis, isolated from the haemolymph of tsetse, were cultured on a layer of Aedes albopictus feeder cells (Welburn et al., 1987). Since Sodalis exists both extracellularly in the haemolymph and intracellularly within tsetse midgut epithelial cells and other tissues, the invasive form may represent a pathogenic remnant. Analysis of the Sodalis transcriptome indicates that quorum sensing may be used to alter gene expression in response to intracellular symbiont density (Pontes et al., 2008). It is further suggested that this may be a key adaptation

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strategy, with Sodalis and host tissues both modulating gene expression profiles and metabolic activities so that immune responses (in particular oxidative stress responses) are minimized. When the impact of different stresses on all three symbiont communities in the tsetse was assessed, the authors concluded that there was a distinct flexibility in symbiont density (with the exception of the teneral stage of the insect), which may help reduce friction between symbionts and host tissues (Rio et al., 2006). The biological contribution, if any, of Sodalis to the tsetse is unknown. A mutualistic relationship has been postulated (and generally accepted) as functional genes encoding enzymes required for vitamin synthesis (Akman and Aksoy, 2001; Akman et al., 2002) (as observed in Wigglesworthia) are present. Such a facultative association is supported by research showing that tsetse longevity is reduced in flies lacking Sodalis (Dale and Welburn, 2001). In addition, puparia that are heavily infected with Sodalis display increased survival under adverse conditions (Baker et al., 1990). It has been suggested that Sodalis may influence tsetse vector competence. Increased trypanosome midgut infections were reported in both lab-reared and wild tsetse that hosted high densities of rickettsia-like organisms (RLOs) (Maudlin and Ellis, 1985; Maudlin et al., 1990). By comparing genetically susceptible and refractory lines of G. m. morsitans, it was observed that the midguts of the susceptible line were prone to the heaviest symbiont infections. To further investigate the role of symbionts (the involvement of Sodalis is inferred) in vector competency, comparisons between refractory lines of G. m. morsitans (Maudlin, 1982) and G. m. centralis (Moloo and Kutuza, 1988a) showed that parasite susceptibility is a maternally inherited factor. In addition, puparial development at lower temperatures revealed a drastic reduction of RLO numbers in teneral flies. This symbiont-depressed tsetse population subsequently produced flies with a greater resistance to trypanosome infection (Welburn and Maudlin, 1991). Examination of the midguts of wild caught tsetse also revealed a direct correlation between trypanosome infections and the density of RLOs. Indeed, it was stated that a wild fly carrying RLOs was six times more likely to be infected with trypanosomes than a RLO-free fly (Maudlin et al., 1990). However, other populations of wild fly do not reflect such a direct relationship between bacterial densities and tsetse refractoriness or susceptibility (Moloo and Shaw, 1989; Geiger et al., 2005b). The vectorial competence between different species of Glossina is highly variable (Harley and Wilson, 1968; Moloo and Kutuza, 1988a,b; Reifenberg et al., 1997; Kazadi et al., 2000). Using this fact as a basis, Geiger et al. (2005b) demonstrated no correlation between Sodalis prevalence and maturation of a Trypanosoma congolense infection in two distinct species of tsetse. However, they could not rule out a potential role for Sodalis in the initial parasite establishment phase in the fly midgut. Further work from the same group investigated the genetic diversity between Sodalis isolated from the same two species of flies using amplified fragment length polymorphism markers (AFLP) (Geiger et al., 2005a). The data

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(albeit weak) implied that vector competence was related to the genetic diversity of Sodalis (not merely the absence/presence of the symbiont) as the symbiont populations from each fly were distinct. The genetic diversity of Sodalis was further interrogated by screening symbionts isolated from trypanosome infected and uninfected flies (Geiger et al., 2007). The ability of a specific parasite species to establish in the insect midgut is statistically linked to the Sodalis genotype present. To address host-specificity among Sodalis, Weiss et al. (2006) transinfected two species of tsetse (previously cleared of native Sodalis with ampicillin) with reciprocal Sodalis strains. Equal symbiont densities were obtained in surrogate hosts without seriously deleterious effects on the fly. Regrettably, determining the effect of Sodalis transfection on the flies’ ability to vector trypanosomes was not reported. Although the above research does not unequivocally link the tsetse symbionts to vector competence it does suggest that symbionts are involved and forms a strong foundation for future investigations.

4

Towards new methods of disease control

Current interventions for management and control of trypanosomiasis are focused predominantly on drug chemotherapy and tsetse fly control especially for the control of nagana (Torr et al., 2005). While both these existing methods are successful, their implementation over extended time periods has not always been sustainable (Vale 1982; Aksoy et al., 2003). Additionally, concern over the development of resistance to available drugs and their toxicity is mounting, and the prospects for new drugs to treat HAT and nagana are bleak (Legros et al., 2002; Fevre et al., 2006). New means of controlling HAT are urgently required. We may find new ways of controlling trypanosomiasis by exploiting the new information becoming available from the genome projects on both tsetse and trypanosomes. To exploit these resources to their fullest we need a means of studying gene function in tsetse and means of expressing transgenic constructs in the fly. 4.1

GENE KNOCKDOWN IN GLOSSINA

The very slow reproductive rate in tsetse (one offspring every 9 days; 40 days adult to adult) and the high costs of maintaining colonies have prevented maintenance of multiple mutant fly lines. In addition, the unusual, viviparous reproductive system of tsetse flies prevents the development of germline transgenesis. Consequently, gene knockdown through RNA interference (RNAi) (Fire et al., 1998) has been of particular value in the study of tsetse biology and gene function. RNAi operates as a means of gene-specific post-transcriptional knockdown and is triggered by double-stranded RNA (dsRNA) mediated degradation of homologous mRNA sequences in the target organism. In the

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current model of RNAi two major steps are involved. The initiator phase involves recognition of dsRNA (that is expressed in or introduced into the cell) and cleavage into short interfering RNA (siRNA) fragments 21–23 bp in length by the enzyme Dicer (Zamore, 2000; Hammond, 2005). The subsequent effector phase involves the incorporation of cleaved dsRNA into a multi-protein complex, known as the RNA-induced silencing complex (RISC), capable of silencing homologous mRNA transcripts (Siomi and Siomi, 2009). Although several methods exist to administer dsRNA to the organism of interest, these are often very species specific. Stable transgenic expression of RNA hairpin constructs (Tavernarakis et al., 2000; Kennerdell and Carthew, 2000), and the use of recombinant viruses to deliver the dsRNA (Travanty et al., 2004) have been used in some Diptera. However, the most commonly used method for dsRNA delivery to insects is the direct injection of dsRNA into the haemocoel. This was first demonstrated in A. gambiae by Blandin et al. (2002), who showed successful gene knockdown of the HDP defensin. This technique has been successfully extended to the analysis of Glossina genes (Hu and Aksoy, 2006; Lehane et al., 2008; Attardo et al., 2008). To date three G. m. morsitans genes, a transferrin, an attacin and tsetseEP protein, have been implicated in tsetse–trypanosome interactions through use of this technique. Knockdown of these genes by dsRNA injection resulted in a statistically significant increase in trypanosome midgut prevalence (Hu and Aksoy, 2006; Lehane et al., 2008; Haines et al., 2009, submission pending). While it is universally practiced, gene knockdown by direct injection of dsRNA into the tsetse haemocoel (and the physical damage that it causes) is clearly not ideal when studying fly immunity. For example, control injections with PBS result in sustained upregulation of attacin and defensin transcripts in tsetse fat body until 18–30 h post-injection (Hao et al., 2001). Cuticular damage is known to stimulate immune responses in other insects including Bombyx mori and A. gambiae (Brey et al., 1993; Han et al., 1999). Thus any injection based RNAi studies on immunity, particularly those of tsetse–trypanosome interactions, should be interpreted with caution as it is known that the establishment of trypanosomes in the tsetse midgut is influenced by, among other factors, the fly immune system (Hao et al., 2001; Hu and Aksoy, 2006; Lehane et al., 2008). In addition, high mortality rates can occur in flies following injection (Walshe et al., 2009) and this too can complicate the interpretation of experimental results because gene knockdown may have occurred to differing extents in the surviving and killed flies. More recently, successful local gene knockdown in Glossina has been demonstrated by ingestion of dsRNA in a bloodmeal (Walshe et al., 2009). Feeding dsRNA may serve as an alternative and preferable means of delivering dsRNA as oral administration is a more natural route of delivery and less invasive than dsRNA injection. In this study, a comparative analysis of knockdown of the immunoresponsive protein tsetseEP (Haines et al., 2005), at both the transcript and protein level in the midgut, was followed after either feeding or injecting dsRNA. High knockdown efficiency was observed using either method of dsRNA delivery, but a

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significantly lower mortality rate was observed following feeding dsRNA compared to injection. Although the midgut specific tsetseEP protein was successfully knocked down by feeding dsRNA, knockdown of the fat bodyexpressed transferrin gene failed (Walshe et al., 2009). However, this gene can be knocked down by dsRNA injection (Lehane et al., 2008). Failure of the RNAi signal to spread beyond the midgut epithelium may be due to the apparent absence of an ortholog to SID-1, the gene responsible for import of circulating RNAi silencing signals in C. elegans, from available Glossina EST databases (and from the Drosophila genome). So, it is possible that the RNAi signal is not distributed systemically in Diptera (Van Roessel et al., 2002; Winston et al., 2002; Roignant et al., 2003; Dietzl et al., 2007; Jose et al., 2009). Thus, while fed dsRNA can clearly enter midgut cells, it may be unable to cross the midgut epithelial barrier in order to cause gene knockdown in tissues beyond the midgut. If this proves to be the case, it would be a valuable tool in itself because the use of two different methods of dsRNA delivery may permit dissection of tissue-specific gene function by permitting tissue specific gene knockdown. This study was the first demonstration of successful gene knockdown in a dipteran by feeding dsRNA. As gene knockdown by feeding dsRNA has been achieved in two other blood feeding arthropods, Ixodes scapularis and Rhodnius prolixus (Soares et al., 2005; Araujo et al., 2006), it is possible this useful molecular tool may be more widely available in blood feeding insects. 4.2

PARATRANSGENESIS

Transgenesis (i.e. the introduction of foreign genes into a living organism) is currently impossible in tsetse flies due to the viviparous lifecycle. So, there is no prospect of disease control through direct transgenic means. However, one proposed approach to trypanosomiasis control involves the modulation of vector competence through paratransgenesis (Aksoy et al., 2003; Weiss et al., 2008). This involves the genetic transformation of symbiotic bacteria residing in the insect to produce anti-parasitic molecules into the midgut. This approach has proven successful in the laboratory, reducing T. cruzi transmission by R. prolixus via expression of a HDP, cecropin A, by its endosymbiont Rhodococcus rhodnii (Durvasula et al., 1997). The foreign genes expressed by the symbiotic bacterium would have trypanocidal properties that would in principal reduce or eliminate trypanosome midgut establishment, thus breaking the parasite transmission cycle (Durvasula et al., 1997; Aksoy et al., 2003). The endosymbiont Sodalis is the target expression vector in the tsetse fly. This bacterium resides in the tsetse midgut as well as other body sites. Consequently, invading brucei and congolense group trypanosomes would encounter trypanocidal secretion products before the trypanosomes could establish in the midgut. Already, recombinant Sodalis produced in vitro may be reintroduced by microinjection into the mother’s haemolymph and subsequently passed to intrauterine progeny

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in utero (Cheng and Aksoy, 1999). As Sodalis can be cultured in vitro (Matthew et al., 2005) and a genetic transformation system has already been developed (Beard et al., 1993), the potential for successful paratransgenesis is high. To date, several candidate anti-trypanosomal molecules have been identified which could be introduced into Sodalis to control trypanosome midgut establishment. One candidate is tsetse attacin (Hu and Aksoy, 2006). Less obvious, but potentially more powerful candidate molecules are also under investigation. For example, BMAP-18 is a truncated form of the bovine myeloid antimicrobial peptide-27 (BMAP-27). BMAP-27 (Skerlavaj et al., 1996) is expressed by bovine neutrophils and exhibits low toxicity to mammalian cells, insect cells and Sodalis, yet causes rapid death to both BSF and PCF trypanosomes (Haines et al., 2003). The truncated BMAP-18 peptide can kill a variety of kinetoplastid parasites, including trypanosomes and Leishmania, yet exhibits reduced cytotoxicity to Sodalis, mammalian and insect cell lines (Haines et al., 2009). Thus BMAP-18 is also a strong candidate for use in a paratrangenesis approach. Several technical problems will need to be overcome before this technology can be applied to disease control. A suitable biological drive system, such as use of Wolbachia (Aksoy et al., 2003), would be required to ensure replacement of the susceptible wild tsetse population with the refractory population. Additionally, this approach would only be suitable for trypanosome species (brucei and congolense groups) that establish in the fly midgut. Therefore, another tactic would be required to control trypanosome species such as T. vivax, which establish in the mouthparts and would not be affected. How rapidly trypanosome resistance would appear with such a system in place is an unknown and a concern. Therefore, it would probably be advantageous to have a selection of effector molecules available for use in a managed control strategy.

5

Conclusion

This review has come at an important time. The genome resources now available for tsetse flies, their symbionts and the trypanosomes they transmit have dramatically increased possible lines of scientific enquiry. Currently, the GeneDB and Vectorbase data repositories contain comprehensive, annotated Glossina EST libraries and completed annotation of the full tsetse genome is projected for 2011. That will mean the complete repertoire of fly, symbiont and trypanosome genomes will soon be available. There are also a range of molecular tools, including RNAi, proteomics, mutant trypanosome lines and an in vitro culture system for Sodalis, that will further our research endeavours on the enemy within. We believe that the technical resources now available mean that it is time for a more integrated approach in research on tsetse–trypanosome interactions rather than the traditional, more fragmentary, approach based on entomology, bacteriology or trypanosome biology. To this end we have used this review to try to make the pertinent entomological information more available to a wide audience across

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the subject. We have taken an entomological viewpoint to examine the interacting biology of the three major components of the tsetse–trypanosome interaction, the fly, its symbiotic bacteria and the invading trypanosome. We have also tried to highlight the problems we believe are associated with some experimental approaches currently adopted in the entomological field. For example, routinely feeding trypanosomes in a blood meal containing glucosamine; failure to precisely define the starvation period prior to the infective bloodmeal; infecting flies with procyclic rather than blood stream form trypanosomes; the inappropriateness in some studies, particularly susceptibility studies, of using long established fly and trypanosome laboratory cultures with no view to the geographical origins of either. Continued use of these experimental approaches may well create increasing problems in interpretation of data. We believe that more field-based studies, despite their difficulties, would be a valuable addition to the understanding of the tsetse–trypanosome interaction. Also, the majority of studies on this vector–parasite system have centered on T. brucei, with T. congolense and T. vivax interactions comparatively lightly investigated. Given the economic importance of both T. vivax and T. congolense and the available genomic resources more emphasis on relations of these trypanosomes with Glossina is also called for. Finally, we are of the opinion that with major improvements in the technical resources available for experimentation of tsetse–trypanosome interactions the remaining major brake slowing progress in the tsetse–trypanosome research field continues to be the high cost and difficulty of tsetse fly colony maintenance. This difficulty has resulted in only a small number of laboratories studying this phenomenon. It is important that funding bodies are fully aware of this fundamental problem if this field of research is to expand. Acknowledgements The authors thank Alvaro Acosta-Serrano, Leyla Akman, Geoffrey Attardo, Lori Peacock and Terry Pearson for their critical reading of the manuscript and helpful comments. Any remaining errors in the manuscript are the responsibility of the authors. We are also grateful to Ray Wilson for kindly granting permission to use his image of the tsetse fly. References Abubakar, L. U., Bulimo, W. D., Mulaa, F. J. and Osir, E. O. (2006). Molecular characterization of a tsetse fly midgut proteolytic lectin that mediates differentiation of African trypanosomes. Insect Biochem. Mol. Biol. 36, 344–352. Acosta-Serrano, A., Vassella, E., Liniger, M., Renggli, C. K., Brun, R., Roditi, I. and Englund, P. T. (2001). The surface coat of procyclic Trypanosoma brucei: programmed expression and proteolytic cleavage of procyclin in the tsetse fly. Proc. Natl. Acad. Sci. USA 98, 1513–1518.

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