Final report
Area-wide control of buffalo fly and prevention of southward spread using Wolbachia
Project code: B.AHE.0242
Prepared by: Peter James
Queensland Alliance for Agriculture and Food Innovation, University of Queensland
Date published: 30 November 2020
PUBLISHED BY Meat and Livestock Australia Limited PO Box 1961 NORTH SYDNEY NSW 2059
Meat & Livestock Australia acknowledges the matching funds provided by the Australian Government to support the research and development detailed in this publication.
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B.AHE.0242 – Wolbachia control of buffalo fly
Contents Abstract ...... 4
Executive summary ...... 5
1. Background ...... 7
1.1 Buffalo flies ...... 7 1.1.1 Description and biology ...... 7 1.1.2 Economic and welfare effects ...... 7 1.1.3. BF lesions...... 8 1.1.4 Southerly spread of BF ...... 9
1.2 Control of buffalo flies ...... 11 1.2.1 Current controls (and problems) ...... 11 1.2.2 Past control of southerly spread (and problems) ...... 11
1.3 Potential for the use of area-wide approaches against buffalo flies in Australia ...... 12 1.3.1 Chemical-based area wide programmes ...... 12 1.3.2 Autocidal and biologically based approaches ...... 12 1.3.3 Wolbachia and area wide control of buffalo flies ...... 14
2. Methodology ...... 18
2.1 BF colony ...... 18
2.2 Adaptation of Wolbachia in Haematobia cell lines ...... 18 2.2.1 Establishment of cell lines...... 18 2.2.2 Infection of cell lines with wAlbB, w Mel and wMelPop Wolbachia...... 19
2.3 Wolbachia assays ...... 19 2.3.1 Detection and quantification ...... 19 2.3.2 Assay to visualise Wolbachia infection dynamics??...... 20 2.3.3 Wolbachia isolation from HIE-18 cells for microinjection ...... 20
2.4 Transinfection of BF with Wolbachia ...... 20 2.4.1 Microinjection of eggs ...... 20 2.4.2 Microinjection of pupae ...... 21 2.4.3 Microinjection of adults ...... 21
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B.AHE.0242 – Wolbachia control of buffalo fly
2.5 Assessment of fitness effects ...... 21 2.5.1 Survival ...... 21 2.5.2 Adult emergence rate ...... 22 2.5.3 Egg production ...... 22
3. Results and Discussion ...... 22
3.1.1 Haematobia cell line establishment ...... 22
3.1.2 Species confirmation using PCR and chromosomal analysis ...... 22
3.1.3 Transinfection of cell line establishment and maintenance ...... 23 3.1.3.1 Wolbachia replication in HIE‐18 cell lines...... 23
3.2 Microinjection ...... 24 3.2.1 Microinjection of eggs ...... 24 3.2.2 Microinjection of adults ...... 25 3.2.2.1 Growth of Wolbachia in injected females ...... 26 3.2.2.2 Distribution of Wolbachia infection in injected female BF ...... 26 3.2.2.3 Survival of Wolbachia infected females:...... 28 3.3 Microinjection of pupae ...... 29 3.3.1 Wolbachia growth and tissue distribution in pupal –infected buffalo flies ...... 29 3.3.1 Effect of pupal injection with Wolbachia on survival of emergent buffalo flies 31 3.3.2 Effect of pupal injection with Wolbachia on adult emergence ...... 32 3.3.4 Effect of pupal injection with Wolbachia on egg production by buffalo flies ... 32
4 Conclusion ...... 34
4.1 Key findings ...... 36
4.2 Benefits to industry ...... 36
5 Future research and recommendations ...... 37
6 References ...... 37
7 Appendix ...... Error! Bookmark not defined.
7.1 Heading...... Error! Bookmark not defined. 7.1.1 Sub heading ...... Error! Bookmark not defined.
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Abstract Buffalo flies (BF) are the second most costly disease issue in northern Australian cattle causing production loss and welfare impacts and are steadily expanding their range into southern production areas. Modelling indicates the potential for the establishment of sustaining populations of BF as far south as South Australia and south east Western Australia by 2030 under the effects of climate change. Wolbachia are maternally transmitted intracellular bacteria of widespread interest in the development of novel controls for insect pests and disease vectors. Wolbachia could be used in area wide control of BF to collapse overwintering populations, to prevent the spread of Stephanofilaria nematodes associated with the development of buffalo fly lesions or potentially for eradication of regional populations or incursion foci. We have shown that BF is a competent host for Wolbachia and that the three Wolbachia strains tested, originally isolated from mosquitoes and Drosophila, successfully infected and grew in transinfected BF. Furthermore we have developed pupal and adult microinjection techniques that can produce ovarian infections in female BF, essential for spread of the bacteria through BF populations, and shown that Wolbachia caused a reduction in lifespan, reduced fecundity and egg hatch and reduced and delayed pupal emergence in infected flies. We are now well placed to produce Wolbachia-transinfected BF lines suitable for use in future biocontrol programs for BF population suppression or local eradication.
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Executive summary
Buffalo flies (BF), Haematobia irritans exigua, are the second most costly disease issue in northern Australian cattle, causing production loss and welfare impacts, and are rapidly spreading southwards. Modelling indicates the potential for sustaining populations of BF as far south as areas of South Australia and south east Western Australia by 2030 under the effects of climate change.
Currently, control methods for BF are almost exclusively applied on a herd-by-herd basis. In the past legislatively supported regulatory programmes based on compulsory chemical treatments and regulated cattle movements were implemented to prevent southerly incursion but these were not effective and were eventually abandoned. Modelling studies indicate the relative inefficiency of herd-by-herd approaches and suggest that application of control techniques on an area-wide basis, targeting the entire population, can be more efficient. This would appear to be particularly the case with BF which overwinter in protected foci of slowly breeding flies from where they spread out to reinfest when conditions become favourable in summer each year.
Wolbachia are intracellular, insect-infecting bacteria of much current interest for use in novel approaches for the control of insect pests and insect-vectored diseases. Wolbachia spreads from mother to offspring in the eggs, rather than horizontally between insects as with most pathogens. Matings between infected males and non-infected females produce sterile eggs whereas matings of infected females produce fertile, infected eggs and this aids the spread of Wolbachia through insect populations. Wolbachia can also cause a range of fitness effects in infected insects and can block the transmission of secondary parasites and pathogens. Wolbachia could be used against BF in three main strategies; (i) by spreading through BF populations and causing fertility and fitness effects that reduce the viability of BF populations, particularly during the overwintering phase; (ii) by blocking the transmission of Stephanofilaria, a nematode transmitted by BF which is implicated in the development of BF lesions; or (iii) by the release of Wolbachia infected males that produce infertile eggs when they mate to uninfected females. This technique, known as the incompatible insect technique (IIT) could be used to eradicate foci of overwintering populations of buffalo flies to prevent southerly spread or to slow rates of re-colonisation of favourable northern areas in summer.
The objectives of this project were (i) to transinfect laboratory populations of buffalo flies with strains of Wolbachia, (ii) to assess the effect of Wolbachia infection on survival and fitness of buffalo flies and (iii) to evaluate the potential for use of Wolbachia to control BF populations, for example in regional eradication programs or to arrest the southerly spread of BF.
To adapt Wolbachia to the BF cellular context and to provide a source of Wolbachia for microinjection we first established a Haematobia cell line (HIE-18). The identity of the cells was confirmed by PCR and cytology and the conditions for growth optimised before the introduction of Wolbachia. The HIE-18 cell line has now been subcultured through more than 200 passages indicating that the HIE-18 cells have formed a continuous cell line. Cryopreserved and subsequently thawed HIE-18 cells have been found to be viable and mitotically active, indicating that they can be maintained in long term storage.
Cell lines containing 3 strains of Wolbachia, wMel and wMelPop originally isolated from Drosophila melanogaster and wAlbB isolated from Aedes albopictus mosquitoes were sourced from collaborating laboratories for use in the transinfection studies and established in our laboratory. Wolbachia were extracted from the host cells as needed, and after a number of attempts, successfully used to establish stably infected Haematobia cell lines. At the time of preparation of this report the wAlbB infected cells had been passaged 125 times, the wMel line 112 passages and the
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wMelPop 98 passages, suggesting that the three Wolbachia strains are well adapted to the Haematobia context, particularly those from more recent passages.
Embryonic microinjection, directly into the eggs, is the most commonly used method for transinfection into new insect species, but proved to be unsuitable for use with BF. The tough outer chorion of BF eggs caused rapid blunting and breakage of microinjection needles, resulting in significant trauma to the eggs. This resulted in very low hatch rates and despite a range of technique modifications and that more than 7,000 eggs were injected, no second generation infections were detected.
Microinjection into the metathorax of adult flies, using a hand held micromanipulator proved to be a much more successful approach with a low mortality rate of adults and infection rates of more than 95% with all three Wolbachia strains. Using adult flies enabled the injection of greater quantities of Wolbachia, improving the likelihood of infection. In addition, as females could be identified and preferentially chosen for injection, this doubled the efficiency of injection compared to eggs and pupae where approximately 50% of the insects injected were males that represent dead end hosts for Wolbachia. Despite an initial reduction in Wolbachia numbers, probably as the result of the host immune response, the infection recovered and became widespread in body and organ tissues. Although Wolbachia infection was not initially detected in ovarian tissues, infected insects were detected in later generations with all three strains, and up to the 4th generation in one case, indicating that ovarian infections had established in a number of instances. Pupal injection resulted in earlier and higher levels of somatic and organ infections as well as ovarian detections in a number of instances. This was considered likely to increase the frequency of trans generation (“vertical”) transmission
Investigation of the effects of Wolbachia on fitness traits in infected flies indicated reduced longevity, decreased and delayed adult emergence, and reduced fecundity in the infected flies, suggesting potential for use in BF suppression strategies. Effects were greatest with wMelPop and wMel and least with wAlbB, which reflected rates of growth of Wolbachia seen in both the cells and infected flies.
Most current BF control programs rely on direct animal treatments. These measures are expensive, often labour intensive and when chemical insecticides are used, risks the development of resistance, occupational health and safety exposure, residues in produce and environmental contamination. The approach suggested here, directly targeting pest populations rather than using individual animal treatments would provide significant labour savings on-farm and would enhance the reputation of Australian produce as clean, safe and ethically produced. Further, BF are continuing to extend their range southwards and the impacts of BF are expected to increase under the effects of climate change. A Wolbachia-based approach offers a novel, convenient and practicable approach to arresting the range expansion of buffalo flies where chemical treatments and regulated cattle movements have previously proven unsuccessful, as well as a pathway to area-wide suppression or elimination of BF populations.
We have now developed the key techniques and critical resources required for the successful development of a Wolbachia based buffalo fly control approach and despite that to date we have conducted relatively few attempts using pupal and adult microinjection, we have now achieved ovarian infections and transmission across generations on a number of occasions. Using these methods and together with the rapid advances in Wolbachia technology currently being made in other labs we believe that there is a high probability that further work in this area will lead to the establishment of Wolbachia-infected BF that can form the basis for novel area wide controls.
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1. Background
1.1 Buffalo flies
Buffalo flies (BF) are exotic pests introduced to Australia from South East Asia in 1938. From there they have spread through most of the northern Australian cattle producing areas and are continuing to spread southward along the eastern coast. Buffalo flies are obligate parasites of cattle and also breeding on buffalo when they are present They are extremely closely related to horn flies, a similar invasive pest in the Americas, which was introduced to eastern USA from Europe in 1886, rapidly spread through north America and has more recently spread through most of the cattle production areas of South America. Both the male and the female subsist completely on blood, using their sharp mouthparts to pierce the animal's skin, causing production loss and irritating cattle. They also cause lesions on cattle, most common near the eyes but also prevalent on the neck, dewlap, belly and flanks that can range from dry and hairless or scab encrusted areas to open and suppurating sores.
1.1.1 Description and biology
Buffalo flies (BF) (Haematobia irritans exigua) are small grey biting flies, about 3-5 -4 mm with two well defined stripes on the thorax. They breed exclusively on cattle or buffalo and although they also bite horses and dogs and sometimes people, they cannot breed on these hosts. If uncontrolled, infestations may reach several thousand flies per animal, each feeding up to 40 times daily, irritating cattle and causing production loss and welfare impacts. The eggs are laid in freshly deposited dung and females ready to lay eggs will concentrate on lower parts of the body until the animal defaecates. Then the female fly will leave the host, deposit her eggs on or beneath the freshly dropped dung and return to a cattle host as soon as she has completed laying. The eggs hatch in 15- 24 h and the maggots feed and go through larval stages in approximately 4-5 days before forming a pupae in the soil. The time to complete these stages is dependent on temperature taking as little as 12 days in summer, but much longer in winter and is also dependent on moisture so that populations build up most quickly when it is warm and wet (Cook and Spain 1982). Buffalo flies usually die out in winter in areas as temperatures drop and conditions become drier and usually can’t survive in areas subject to frosts. BF do not have a particular overwintering stage, as present in horn flies, but rather populations contract to very favourable areas and populations survive in these foci some as low numbers of slowly cycling flies, in warmer moist protected areas. When temperatures begin to warm and the rains come fly populations build and spread out from these foci by movement with cattle and by autonomous flight.
1.1.2 Economic and welfare effects
BF are estimated to cost the beef industry $99m p.a. in an average year and are second only to ticks in terms of annual economic cost of health issues in northern cattle herds (Lane et al. 2015). Clearly costs are much higher than this in severe BF years. BF were ranked first by northern beef producers in a survey of animal health problems in 1992 (Bock et al. 1995) and when dairy farmers were surveyed in 1998, 61% said BF were a significant problem and 29% said they were a small problem whereas for ticks the corresponding figures were 13% and 43% (Jonsson and Matchoss 1998).
Estimated reductions in growth caused by BF infestation have varied greatly, often because of difficulties in developing a robust experimental design. Bean et al. (1987) found that animals treated for BF gained weight 15%, 11% and 4% in the three years of their experiment and Spradbery and Tozer (1996) found that cattle treated with insecticidal ear tags gained 13 kg more body weight than
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untreated cattle over a 5 month period. Holroyd et al (1984) reported significantly improved weight gains from treatment in 2 of 4 years, but noted that insecticide application in the treatment groups may have affected fly numbers in the controls and that a positive correlation between body weight and fly numbers may have obscured the beneficial effects of treatment. Results correspond approximately with those from studies with horn fly (Haematobia irritans irritans; HF) in North America where improvements in weight gain of 8.3% (Kinzer et al 1984), 8.7% (Harvey and Brethour 1979), 11% (Harvey and Brethour 1981) and 18% (Haufe 1982) were recorded.
Effects on skin quality have not been quantified in Australia. Lesions penetrate the dermis and render the affected areas unusable (Sutherst et al. 2006). They can cause significant wastage if they occur in valuable areas of the hide. Guglieomone (1999) describes an increase in hide damage noted by tanneries that coincided with the spread of HF through Argentina and Uruguay. Up to 50% of the hide was affected by black spots, pitted areas and semi-circular scars resulting from HF feeding. Similar effects are likely to contribute significantly to lower values received for hides from northern cattle (Sackett et al. 2006) and could be expected to cause very significant reductions in skin values of more susceptible southern cattle.
The estimates above did not include animal welfare impacts, thought to be considerable in a number of quarters. A survey of dairy farmers in the tick infested areas of Qld in 1997 listed buffalo flies as a key animal health problem (Jonsson and Matchoss 1998). BFwere considered to be a greater concern than ticks with 55% considering them a significant problem, compared to 14% for ticks, and only 6% considering them not to be a problem, compared to 32% for ticks. When asked what aspect of BF infestation worried them most, 40% said production loss and 42% said the welfare impacts. Control of BF is increasingly compromised by the development of resistance to control products and there are rapidly growing markets for meat and milk produced in low- chemical systems. The move towards a greater proportion of Bos taurus genes in northern cattle herds to raise beef quality is expected to increase susceptibility to BF.
1.1.3. BF lesions
BF feeding can lead to the development of lesions that are of welfare concern (Jonsson and Matchoss 1998), reduce hide value and make cattle less acceptable for the market (Lane et al. 2015; Guglielmone et al 1999). In addition, the lesions present a potential focus for strikes by Old World screwworm Chrysomya bezziana (Villeneuve) flies, which are endemic in a number of Australia’s nearest northern neighbouring countries and which are considered a major biosecurity risk for northern Australia (Animal Health Australia 2017). These lesions can range from dry, alopecic, or scab encrusted to severe open and suppurating (Johnson 1989). They are found most commonly beneath the eyes of cattle, but can also be prevalent on the neck, dewlap, belly and flanks (Sutherst et al. 2006) and their development and persistence has been associated with a currently unnamed species of filarial nematode (Stephanofilaria sp.), vectored by BF (Johnson 1986; Shaw and Sutherland 2006).
Lesions are most widespread in northern areas of Australia, where BF are present throughout the year, with up to 95% of cattle affected (Johnson et al. 1986). It is expected that the prevalence of lesions will increase in more southern parts of the fly range as global warming extends the length of the BF season and the intensity of fly attack.
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1.1.4 Southerly spread of BF
1.1.4.1 Historic spread of buffalo flies
BF entered mainland Australia near Darwin (12.5° S, 130.8° E) in 1838, probably on water buffaloes (Bubalus bubalis L. 1758) introduced from Timor in 1825 (Tillyard 1931). Early spread occurred very slowly and coincided closely with the spread of buffaloes (Hill 1917), which appear to be the preferred native host of the flies in Asia (Iwasa and Ishigura 2010) (Fig. 1).
Figure 1: Spread of BF in Australia.
It wasn’t until 1928 that buffalo flies reached the Queensland border, approximately 1300 km south- west of their original point of introduction (Seddon 1967), subsequently spreading across the dry stretch of land south of the Gulf of Carpentaria to eastern Queensland during a series of wet years in 1939-41. From there, they spread rapidly to the east coast of Cape York in northern Australia and southwards along the eastern coast until they appeared to reach a southerly limit just north of Bundaberg (24.8°S latitude) by 1946. Here the spread paused, and no further southerly spread was observed for the next 30 years (Figure 1).
Following a series of mild winters and wet years from 1973 onwards, changes to BF and cattle tick (Rhipicephalus australis) regulatory programmes, and possibly aided by changes in the chemicals used for cattle tick treatments, southerly range expansion recommenced and BF reached the Brisbane Valley and Nambour in 1977, the Tweed Valley in New South Wales in April 1978, and Bonville, south of Coffs harbour (30.4°S) in 1982 (Williams et al. 1985). Since then, the flies have continued their southerly spread with infestations seen as far south as Dubbo, Narromine and Maitland (32.7°S) in 2011. This represents an increase in their southerly range of approximately 1000 km in the last 40 years. Following their first detection in New South Wales in 1978, the flies now
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survive the winter in many eastern parts of the state and have become a significant endemic cattle pest in these areas.
The impact of BF and the area affected in Australia varies significantly with season and weather conditions. During warm wet summers the distribution of the flies increases significantly in northern and north-eastern areas, and they may spread to affect cattle in an area potentially more than two times larger than the permanently infested range (Fig. 1).
1.1.4.2 Effects of Climate Change
All indications to date suggest that climate change effects in Australia will facilitate the continued spread of BF into new areas and will increase the economic and welfare impacts in the southern parts of their current range (CSIRO/BOM 2016). Rising temperatures will enable more rapid population growth, an increased number of generations each year, greater fly activity in many areas and longer seasons of cattle challenge. In addition, predicted rises in minimum temperatures and a reduction in the frequency of frosts will favour survival in marginal areas and further southerly extension of the flies’ range. A possible increase in the summer incidence of rainfall in some areas of Australia may also favour the flies’ breeding.
The results of CLIMEX modelling (R. Dobson personal communication 2015) suggest greater impacts from BF in the southern parts of their current range, including the potential for persisting fly populations to establish through most of the moist coastal belt of New South Wales and in foci as far south as South Australia and southern Western Australia (Fig. 2). In addition, increased weather variability and extreme rainfall events predicted under climate change may assist the spread of flies across inhospitable areas to new foci suitable for winter fly survival. Once established in these areas, new overwintering foci would provide a source for more extensive incursions during warm wet periods, similar to that seen in northern Australia.
The CLIMEX modelling does not account for factors such as a changing resource base, microclimate effects or changes in pest biology. In southern areas, the cattle industry is based largely on Bos taurus L. breeds that are more susceptible to BF than the Bos indicus L. cattle that predominate in northern areas (Frisch et al. 2000). In addition, northern cattle are normally treated to control cattle ticks, which can also impact on BF numbers, whereas few parasite treatments are applied to southern cattle. Thus, the southern beef and dairy industries provide a susceptible and largely untreated host resource extremely favourable for invasion by BF. In addition, adaptation of insects at the edge of their range can be an important contributing factor in new pest invasions (Hill et al. 2011).
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Predicted 2030
Figure 2: CLIMEX predictions of areas suitable for the establishment of BF populations by 2030 with predicted climate change. Solid circles indicate favourability for BF, crosses indicate weather station sites unsuitable for BF persistence.
The degree to which genetic adaptation of BF to cooler temperatures has contributed to their southerly spread is uncertain. However, Iwasa and Ishigura (2010) note that the flies in their native range appear to prefer buffalo to cattle as hosts and a period of adaptation to cattle and cooler Australian conditions may have contributed to their spread in Australia. Development of pupal overwintering capacity as they move south, which must be considered a possibility given their close genetic relatedness with HF, is a concerning prospect and could see the species develop a temperate distribution in Australia, similar to that seen for HF in the northern hemisphere and South America.
1.2 Control of buffalo flies
1.2.1 Current controls (and problems)
Currently control of BF in Australia depends largely on chemical treatments, although techniques such as BF traps (Sutherst and Tozer 1995) and selection of Bos indicus breeds (Frisch et al. 2000) are also used. In addition, dung beetles may assist the regulation of BF populations under some circumstances (Doube 1986).
1.2.2 Past control of southerly spread (and problems)
At various stages in the spread of BF in Australia, regulatory programmes, supported by legislation, and which included movement controls and compulsory spraying of relocated cattle with insecticides, were used in an attempt to prevent their southerly incursion (Parliament of Queensland 1965). However, these programmes were not effective in stemming the southward spread of BF (Anonymous 1934; Roberts 1946; Eastaway 1974) and all were eventually abandoned in 1978 (Williams et al. 1985). However, BF remain a legislatively specified notifiable disease in some southern states of Australia, where the flies are not currently present (Department of Agriculture and Water Resources Australia, 2017).
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1.3 Potential for the use of area-wide approaches against BF in Australia
Currently, control methods for BF are applied almost exclusively on a herd-by-herd basis. However, modelling studies indicate the likely inefficiency of herd-by-herd approaches showing that the effects of invasion of pests from untreated areas can be devastating in compromising the effectiveness of control measures (Knipling 1972a). Application of control techniques on an area- wide basis, targeting the entire population rather than just individual properties or herds, can be much more efficient than more intensive programmes applied on a herd-by-herd basis. Area-wide approaches are expected to be particularly advantageous when pests are mobile and can readily auto-disseminate and therefore may not be easily controlled by property-based or herd-based programmes (Hendrichs et al. 2007), such as is the case with BF.
1.3.1 Chemical-based area wide programmes
Area-wide control programmes have historically been based mainly on the application of chemical pesticides by methods such as aerial spraying or intensive ground spraying, or in the case of diseases of livestock, by individual animal or herd treatments with quarantine controls and movement restrictions (Graham and Hourigan 1977). At various stages in the spread of BF in Australia, regulatory programmes, supported by legislation, and which included movement controls and compulsory spraying of relocated cattle with insecticides, were used in an attempt to prevent their southerly incursion (Parliament of Queensland 1965). However, these programmes were not effective in stemming the southward spread of BF (Anonymous 1934; Roberts 1946; Eastaway 1974) and all were eventually abandoned in 1978 (Williams et al. 1985). However, BF remain a legislatively specified notifiable disease in some southern states of Australia, where the flies are not currently present (DAWR 2017).
1.3.2 Autocidal and biologically based approaches
Programmes which require widespread application of pesticides are increasingly unacceptable on a community basis and can be compromised by the development of resistance or resurgence of pests from cattle that are not treated or where treatments are poorly applied. More biologically-based, species-specific and environment-friendly techniques which operate by disrupting biological processes of pests, generally find wider community acceptance and are often more effective than pesticide applications (Bourtzis et al. 2016). In addition, because of the ability of released insects to disperse into all areas occupied by the target field population and to actively search out and mate with target insects, biologically-based methods are often more effective against pests that can disperse autonomously or which survive in cryptic habitats that are hard to reach with chemical sprays.
The most well-known of these approaches is the sterile insect technique (SIT) in which insects of the target species are mass-reared and sterilised using low level ionizing radiation and then inundative releases of the sterilised insects (usually males) are made on an area-wide basis over the entire area of the target population (Dyck et al. 2005; Vreysen and Robinson 2011). The sterile males mate with field females, which consequently produce infertile eggs and through sequential releases the target population is suppressed, or under certain conditions, eradicated.
Some of the most significant successes with SIT have involved insect pests of livestock, including the eradication of New World screwworm fly (Cochliomyia hominivorax (Coquerel); NWSF) from North and Central America (Wyss 2000), eradication of an incursion of NWSF in Libya (Lindquist et al.
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1992), and removal of the tsetse fly Glossina austeni (Newstead) from the Island of Unguja in the Zanzibar archipelago (Vreysen et al. 2000; Vreysen et al. 2014). Localised eradication or suppression using SIT has also been achieved on a number of occasions with other tsetse species in Africa (Vreysen et al. 2013). Successful eradication using this approach can be extremely cost efficient. For example, in the case of the NWSF programme in the Americas it has been calculated that the direct benefits achieved each year from the programme are equal to or greater than the total cost of the sterile male release programme over the fifty years of its operation (Vreysen and Robinson 2011).
HF, which as noted are very closedly related to BF, are one of the species suggested by Knipling (1972b) as likely candidates for control by SIT. Knipling considered that the close association of the flies with cattle and its consequent accessibility to control meant that fly populations could be readily reduced by insecticide treatment of cattle, then the remaining population eliminated using SIT. In early trials with HF, cattle were sprayed with topical pesticides to reduce fly numbers. However, the subsequent sterile insect releases were compromised because the released flies were more susceptible to the insecticides used than were the field flies (Eschle et al. 1973, 1977). This was overcome by using methoprene, an insect growth regulator administered in drinking water, which targeted the larval stages of HF, and had no effect on the released adult flies. Trials on the isolated Kalaupapa peninsula of Molokai in Hawaii subsequently confirmed that a semi-isolated population could be effectively eradicated using this method, even in the very HF-favourable environment of Hawaii (Eschle et al. 1977). Unfortunately, the area was later reinfested by the introduction of infested cattle into the area.
Although the SIT is by far the most widely known and successful genetic technique used against livestock pests to date, SIT is not always feasible and a range of other genetically-based techniques have also been tested, or are under contemplation. For example, in continental Australia the extensive areas of livestock production and the wide distribution of associated pest species, together with few natural geographic boundaries, made the use of SIT impractical or at least of dubious cost-benefit for use against many livestock pests as in the case of sheep blowfly Lucilia cuprina (Wiedemann). As a consequence, a range of other genetically-based techniques such as the use of compound chromosome and sex-linked translocation strains (field female killing systems), which were predicted to be more effective at lower release ratios, were developed and tested (Foster et al. 1985, 1988, 1991). Field testing showed promise for these approaches, but for a number of reasons discussed by Scott (2014), they were never implemented for widespread use.
More recently, transgenic sexing strains of L. cuprina have been developed that carry a tetracycline- repressible female lethal genetic system that could form the basis for mass-production of only males of L. cuprina, and potentially other fly species, for use in genetic control programmes (Scott 2014). A range of other techniques such as the release of insects carrying dominant lethal genes (RIDL), RNAi and homing endonuclease genes (HEG) are now also being considered for use with mosquitoes, tsetse flies and other species (reviewed by McGraw and O’Neill 2013; Bourtzis et al. 2016) and have proceeded to field testing in some instances (Harris et al. 2011). With increasing access to sequenced insect pest genomes (International Glossina Genome Initiative 2014; Anstead et al. 2015) and rapid advances in molecular technology, most notably the availability of new gene editing techniques such as CRISPR/Cas9, many new, purpose-designed approaches for control will likely emerge.
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1.3.3 Wolbachia and area wide control of BF
Of much current interest for use in area-wide control programmes are symbiont-based approaches (Bourtzis 2008; Bourtzis et al. 2016; McGraw and O’Neill 2013; Wilke and Marelli 2015), in particular the use of Wolbachia. Wolbachia are maternally transmitted intracellular bacteria in the family Alphabacteria estimated to infect 40% of terrestrial arthropod species (Zug and Hammerstein 2012). Wolbachia are capable of spreading through insect populations by manipulating host reproductive processes and have many and varied other effects that present potential for use in BF control programmes (Hoffmann et al. 2015). These can be considered in three main groups, a) cytoplasmic incompatibility, which can be harnessed for population suppression, population replacement or potentially population elimination; b) fitness effects induced by Wolbachia infection; and c) transmission blocking of secondary pathogens. These strategies are considered below for their potential to reduce the impacts of BF or interrupt their spread into uninfected areas.
1.3.3.1 Cytoplasmic Incompatibility and Incompatible Insect Technique (IIT) Wolbachia infection can interfere with insect reproduction in a number of ways, including through the induction of cytoplasmic incompatibility whereby matings between infected males and non- infected females or between males and females infected with incompatible Wolbachia strains (bidirectional incompatibility), produce infertile eggs. This approach when used as an insect suppression or eradication strategy has been termed the incompatible insect technique (IIT) (Zabalou et al. 2009). The IIT method is similar in approach to SIT, with Wolbachia-infected males used as de facto sterile males. Since Wolbachia is not paternally transmitted, as long as similarly infected females are not also released, the Wolbachia strain present in the released males does not establish in the field. Serial release of the infected males can lead to population suppression or eradication.
The effectiveness of using Wolbachia-induced cytoplasmic incompatibility in this way was demonstrated as early as the 1960s when release of Wolbachia-infected male Culex quinquefasciatus Say mosquitoes, vectors of human filariasis, led to local eradication of this species from areas in Myanmar (Laven 1967). Since then studies towards the use of IIT have been conducted with a range of mosquito species, including Aedes polynesiensis Marks (Brelsfoard et al. 2009; O’Connor et al. 2012), Aedes albopictus (Skuse) (Calvitti et al. 2010), Anopheles stephensi Liston (Bian et al. 2013) and Culex pipiens pallens (Coquillett) (Chen et al. 2013), and the veterinary pests Glossina morsitans Westwood (Alam et al 2011; Bourtzis et al. 2016) and Stomoxys calcitrans (L.) (Kusmintarsih 2009).
Use of an IIT approach could be applicable for eradication of confined foci of overwintering populations of BF to prevent or retard southerly spread or to slow rates of re-colonisation of favourable northern areas in summer. The IIT method could also be used to eradicate BF that become established in relatively isolated areas as a result of climate change, such as those predicted in South Australia and south-western Western Australia (Fig. 2).
Ideally only male BF would be released, but to date no method for accurate mass-sexing of HF or BF has been reported. In the case of the Hawaii SIT trials with horn flies, irradiated flies of both sexes were released (Eschle et al. 1977). Although this is usually undesirable, because it increases competition with field females for mates and can temporarily increase fly pressure on cattle, it did not compromise success in the case of the Hawaiian trial and may not be a consideration if used against low-level populations of flies present in overwintering foci of BF.
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Reduction of male mating competitiveness from the effects of irradiation is one of the difficulties sometimes experienced in SIT programmes (Zhang et al. 2015). As female flies are often sterilised at levels of radiation below that which causes reduction of competitiveness in males, this has led to the suggestion of the complementary use of SIT and IIT, with Wolbachia used to induce functional sterility in the males and low-level irradiation used to sterilise the females (Brelsfoard et al. 2009; Zhang et al. 2015; Bourtzis et al. 2016). In the absence of a practical sexing method, a similar approach could be considered for BF. Alternatively, the development of a self-sexing strain in stable flies S. calcitrans (Seawright et al. 1986), which are in the same subfamily as BF, the determination of a Near Infra Red (NIR)-based method for sexing tsetse fly pupae (Dowell et al. 2005), and the rapid advances with molecular techniques currently being made in other species (Scott 2014), suggest significant potential for the future development of a sexing method for BF. Notwithstanding the potential added difficulties for artificial rearing, the use of a strain of Wolbachia that also confers a fitness disadvantage or inability to overwinter in infected flies, such as wMelPop (see below), is a further possibility to guard against the effects of inadvertent female release in a Wolbachia-based IIT programme.
1.3.3.2 Using Wolbachia-induced fitness effects to collapse overwintering Populations of BF
Different strains of Wolbachia can induce a range of different effects on the fitness of infected hosts (Hoffmann et al. 2015). Some of these effects include reduced lifespan (McMeniman et al. 2009), mortality of eggs (McMeniman and O’Neill 2010), slowed larval development (Ross et al. 2014), and reduced overall fitness (Yeap et al. 2011, 2014; Ross et al. 2015). Infection with Wolbachia has also been shown to interfere with blood feeding efficiency in mosquitoes (Moreira et al. 2009; Turley et al. 2009), and to affect locomotor activity in parasitic wasps, Drosophila species, and some mosquitoes (Fleury et al. 2000; Peng et al. 2008; Evans et al. 2009). Similar effects in BF could also have deleterious effects on survival and mating efficiency, as well as the persistence of BF populations, particularly during winter.
The most profound deleterious effects described have been from the ‘popcorn’ (wMelPop) strain of Wolbachia, initially isolated from laboratory populations of Drosophila melanogaster (Min and Benzer 1997). The wMelPop strain replicates in host cells, causing cellular damage, characteristic morphological changes in infected tissues, and a range of physiological effects. These effects reduce lifespan by approximately one-half in D. melanogaster and transinfected mosquitoes (Min and Benzer 1997; McMeniman et al. 2009). Reductions of lifespan of this magnitude, and other fitness characters, can have profound effects on the population dynamics of a species, particularly during unfavourable times of the year (Rasic et al. 2014). However, the effects of Wolbachia are highly strain-, host- and environment-dependent, and less profound effects on fitness have also been observed in other Wolbachia-host associations (Hoffmann et al. 2015).
Modelling conducted by Rasic et al. (2014) demonstrated potential for using fitness reductions induced by Wolbachia to suppress or eliminate Aedes aegypti populations, particularly in locally or seasonally variable environments. Their results suggested that the effects of wMelPop were not sufficient to reduce persistence of mosquito populations in the very favourable climates of north Queensland, but were likely to cause local extinctions in the more mosquito-marginal environments of central Queensland. These predictions were supported by semi field cage studies, which showed that reductions in the survival of desiccation-resistant eggs resulting from wMelPop infection, eliminated populations of Ae. aegypti during extended dry periods (Ritchie et al. 2015).
Wolbachia could also be used to drive co-inherited deleterious ‘payload genes’ in the genome of infected insects into the target pest population (Curtis and Sinkins 1998; Hoffmann and Turelli 2013;
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Champer et al. 2016). These genes could confer reduced fitness or conditionally lethal effects such as cold temperature sensitivity or insecticide susceptibility.
Conversely the use of linked traits that confer a fitness advantage in certain circumstances might be used to facilitate the spread of Wolbachia strains into a population. For example, insecticide resistance that confers a competitive advantage under a spraying regime could be used to assist the spread of a wMelPop-infected strain that confers a pathogen blocking capability or seasonal lethality (Hoffmann and Turelli 2013).
As more pest insect genomes are characterised and with rapid advancement in molecular transformation technologies, it is expected that possibilities for this approach will grow rapidly. Using Wolbachia as the driving mechanism is expected to have greater public acceptance and less potential for unanticipated effects than transgenic gene drives (Champer et al. 2016). An attractive alternative approach is the direct transformation of Wolbachia genomes with genes to be driven into a pest population. Until recently, successful genetic transformation of Wolbachia had proved elusive, but the recent reporting of a phage-mediated system for the genetic modification of Wolbachia (Bordenstein and Bordenstein 2017) offers exciting possibilities in this area.
1.3.3.3 Stephanofilaria Blocking
BF-associated lesions are of significant welfare and economic concern, with estimates of over 95% of cattle affected in northern areas of Australia (Johnson 1989). Although the exact aetiology of BF-associated lesions is unclear, an unnamed species of filarial nematode (Stephanofilaria sp.), vectored by BF and found in the lesions, is thought to play a role (Johnson et al, 1986, Johnson 1989). Surveys of BF collected from near Townsville in the 1980s found a prevalence of Stephanofilaria in female flies of 2.9% (range 0% - 9.3%) (Johnson 1989), whereas a more recent study in 2004 measured infection rates between 29% and 57% in flies collected from four sites near Rockhampton (Shaw and Sutherland 2006).
Wolbachia infection has been demonstrated to affect vectorial capacity of mosquitoes for a range of pathogens, including filarial nematodes. Inhibition of development of filarial nematodes was seen with both wMelPop in Ae. aegypti (Kambris et al. 2009) and wAlbB in Ae. polynesiensis (Andrews et al. 2012) and resulted in a reduction in the prevalence of infective third stage nematodes in the mosquitoes. The mechanism of pathogen blocking is not completely understood, but may be due to competition for host resources or modulation of host immune response, in particular reduction in levels of reactive oxygen species (Andrews et al. 2012). The wMelPop strain of Wolbachia also reduces the efficiency of disease transmission by shortening the life span of vectors and reducing the likelihood that a pathogen will be able to complete its required extrinsic incubation period before host mortality. The shortest incubation period seen for Stephanoflaria sp. in BF was 7 days (Johnson 1989), suggesting that life-shortening effects of wMelPop Wolbachia could also significantly affect the transmission dynamics of this species.
Lesions associated with HF-transmitted Stephanofilaria stilesi in North America appear to be less extensive and severe than BF-associated lesions in Australia (Hibler 1966). As HF are infected with Wolbachia, but BF are not, it is tempting to hypothesise that this difference may be associated with differences in the efficiency of Stephanofilaria transmission, although many other factors could also be involved. Disruption of the spread of Stephanofilaria or reduction in the severity of lesions by the introduction of a transmission-blocking Wolbachia strain into BF, would be a significant outcome for the Australian cattle industries from both economic and welfare perspectives.
1.3.3.4 BF overwintering, a susceptible stage for area-wide approaches?
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HF have the ability to overwinter in the pupal phase, as pharate adults, whereas BF do not (Ferrar et al. 1969; Cook and Spain 1982), which is a major difference between the two species (Showler et al. 2014). In the northerly part of their range in North America, adult HF begin to disappear from cattle in autumn and do not reappear until the next spring. Overwintering dormancy allows HF to emerge and rapidly re-establish throughout the previous season’s range when conditions become suitable in spring or summer. There is however, significant plasticity in this response and at warmer latitudes HF populations continue cycling throughout the year (Showler et al. 2014). In more marginal areas HF may survive winter both as adults, with reduced activity, and in the soil stages, with various levels of dormancy. Mendes and Linares (1999) working in a warm winter climate in Brazil (21°30’S), verified diapause in 9.1% of winter pupae, even though HF were present on cattle year-round. These authors note that this dual overwintering mechanism could present difficulties for the design of cost efficient eradication programmes for HF. The plasticity in overwintering response has probably been a key factor allowing HF to disperse and become established in a wide range of environments.
In contrast to HF, BF die out through much of their summer range in winter (Fig. 1). Their range at the southern and continental edges in Australia is limited by cooler temperatures and low moisture levels in dung during winter (Cook and Spain 1982). Low temperatures either prevent development completely, or slow the development of the larval stages to a degree that they can’t be completed before moisture content in dung falls to lethal levels. The occurrence of frosts can also have a devastating effect on the survival of the soil stages (Cook and Spain 1982).
Williams et al. (1985) found that BF overwintered at the edge of their winter range as slowly cycling, low level fly populations in local areas of moderate microclimates. Most of these overwintering foci were in hilly, heavily timbered areas that were well-watered from either creeks, dams or swamps, and less exposed to low minimum temperatures or frosts than the low-lying surrounding areas. Nearly all of the overwintering sites identified were within 40 km of the coast where temperatures were likely moderated by coastal influences. Re-colonisation of summer-suitable areas and southern range extension relied on overwintering of BF in these foci. When conditions became favourable each year the flies built-up in numbers and either dispersed from these areas autonomously or were transported by cattle movements to re-infest their summer range (Fig. 2).
These localised overwintering foci provide a potential target for the application of Wolbachia-based approaches. The use of Wolbachia in either an IIT approach, to compromise Stephanofilaria transmission or to introduce a deleterious fitness factor, is likely to be most efficiently achieved at times of low fly populations, such as during overwintering, when suitable release ratios will be most readily achieved. Indeed, SIT and IIT approaches are often initiated when target populations are low, or involve population reduction by insecticide treatments prior to the release of infected flies. Persistence of BF populations in overwintering foci is precarious and it is the soil stages that are most subject to adverse effects from low temperatures and dryness. Adult flies living with the warmth and blood provided by their cattle hosts are less affected by adverse winter conditions. Therefore, it is likely that released adult flies will be less exposed to the effects of winter conditions than soil stages and able to persist for sufficient time to mate with overwintering adult flies and either interrupt reproduction or spread Wolbachia infection. Objectives
• To transfect laboratory populations of BF with strains of Wolbachia • To assess the effect of presence of Wolbachia on reproduction and survival/fitness traits of BF and model the likely effects on BF populations and economic impact.
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• To evaluate the potential for use of Wolbachia to control BF populations, for use in regional eradication programs and to arrest the southerly spread of BF and to recommend strategies towards practical implementation
2. Methodology
2.1 BF colony
All BF adults and earlier stages used in this project were obtained from a laboratory colony of BF maintained in our laboratory for more than 180 generations. The establishment of and maintenance of this colony were previously described in the final report of project B.NBP.0488. Briefly, for this project, adult flies were housed in 30 x 30 x 45cm aluminium framed cages, with glass ends, mesh sides and cloth sleeve entrance and held in a constant environment room at 28°C and 70%RH, with a 15 hour light and 9 hour dark light regime. Prior to each use cages were disinfected and bottom of the cage covered by Wettex® sheet overlaid with blotting and saturated with tap water top provide extra humidity in the cage. For egging field collected and frozen-stored manure was thawed, fashioned into small artificial pats and placed onto the paper towelling in the cage when egg laying commenced (approx. 5 days old). For eggs collected over one night, no manure was used. For collection over two or more nights, 500 ml quantities of manure were fashioned into artificial pats and placed on the bottom of the cage and kept moist. Blood was provided at the top of the cage from two silicon membrane blood feeders water jacketed and with water heated to 36°C circulated through them from a water bath. The silicone membranes through which the flies fed were c. 0.06mm thickness supported by polyester mesh. When not transinfected the adult BF were fed bovine blood, containing 4.5 g sodium citrate and 50mg/L kanamycin sulfate or, more recently, 0.3ml/L formalin. With flies to be used for transinfection or potentially already transinfected with Wolbachia the formalin was omitted. Flies were also provided ad lib with a vitamin/mineral supplement prepared by dissolving 1 tablet of Hair Skin and Nails Vitamin and Mineral Supplement (Herron Pharmaceuticals) in 500 ml Mount Franklin water delivered from an inverted 200 ml container at the top of each cage.
Eggs for microinjection or from transinfected flies were collected from the filter paper at the bottom of the cage daily, whereas eggs for colony maintenance were collected from the bottom of the cage, or as larvae in the dung and transferred to larger dung pats with moisture adjusted to 80-85% and on sand and allowed to complete development to pupae. Eggs for microinjection with Wolbachia were collected within 10 hours of deposition. Eggs from transinfected flies were collected for testing or reared through to pupation of smaller pats and emergent flies transferred to smaller cages for breeding or use in studies of fitness effects.
2.2 Adaptation of Wolbachia in Haematobia cell lines
2.2.1 Establishment of cell lines
The success rate of microinjection of Wolbachia into new host species is generally low, with subsequent loss of infection in newly injected hosts common, particularly in more distantly related host species. This is thought to be due to inability of the injected Wolbachia to adapt quickly enough to the new host environment. However, it has been indicated that the probability of success can be
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increased by prior adaptation of Wolbachia in more closely related host cell lines (McMeniman et al. 2008, 2009).
Towards this end, four sustaining BF cell lines were established at St Lucia in 2010. Of the four cell lines established, BFEC#12 showed the best growth, was maintained in the laboratory for six months and subcultures were frozen down and stored liquid nitrogen at both the Animal Research Institute at Yeerongpilly and University of Queensland at St Lucia. Half of the BFEC#12 cell lines stored at ARI were retrieved successfully from liquid nitrogen storage and grew vigorously enough for new reserve cell lines to be frozen for liquid N2 storage. Unfortunately, during the Brisbane floods in January 2011, power was cut at both UQ and ARI. Attempts to rescue the cells by addition of conditioned media and/or fly egg extract to the culture were partly successful, such that the cells continued to grow slowly from mid-February. However, bacterial contamination of these revived lines led to them being subsequently discarded. Many subsequent attempts to re-establish this line both in the previous project and the current project have been unsuccessful. However, in a subsequent collaboration with Dr Tim Kurtti and Dr Ulrike Munderloh at the University of Minnesota we were successful in establishing a Haematobia irritans irritans cell line (see Madhav et al. 2020). BF (Haematobia irritans irritans) and HF (Haematobia irritans exigua) are very closely related and generally considered to be subspecies. The larval stages of the two species are extremely similar and are probably morphologically indistinguishable (Pont 1973) and the adults are mainly distinguished morphologically by the presence of 4 to 6 long curled hairs on the hind tarsi of male BF, but even this character is reported to be variable (Snyder 1965). In a study of the COI and COII genes of BF and HF, Iwasa and Ishiguro (2010) found a sequence divergence of only 1-8% to 1.9% between the two species again confirming an extremely close genetic relationship. Therefore to facilitate progress in this project it was decided to move forward using the HIE-18 cell line. Details of the generation of the HIE Cell line and have been previously reported in Final report of MLA project B.NBP.0488 and in Madhav et al. (2020).
2.2.2 Infection of cell lines with wAlbB, w Mel and wMelPop Wolbachia
Infection of the HIE cell lines with wAlbB, wMel and wMelPop Wolbachia was carried out using the protocol described by Herbert and McGraw (2018) after confirming freedom from the wIrr (irritans) strain of Wolbachia previously identified in H. i. irritans (Palavesam et al. 2012; Madhav et al. 2020b) PCR. Briefly, wAlbB‐, wMel‐ and wMelPop‐infected mosquito cell lines were grown in 75 cm2 flasks, each containing 15 mL of M&M medium with 10% FBS. Cells were dislodged from the flask after 7 days by vigorous pipetting and spun at 2000 g. The resulting pellet was washed three times with 5 mL SPG buffer and sonicated on ice at 24% power using a Q125 sonicator (Qsonica, Newtown, CT, USA). Cellular debris was pelleted by spinning at 1000 g for 10 min at 4 °C, and the supernatant was filtered through 50‐ and 2.7‐μm acrodisc syringe filters. Filtered supernatant was centrifuged at 12 000 g, and the Wolbachia pellet was resuspended in 100 μL SPG buffer. The pellet suspension was immediately added drop by drop to the cell line flasks when the HIE‐18 cells were 80% confluent. Infected cell lines were cultured for 5–6 days and then divided between new flasks in a ratio of 1:2. Infected cells were frozen for storage after every tenth passage.
2.3 Wolbachia assays
2.3.1 Detection and quantification
DNA was isolated from cells using an Isolate II Genomic DNA kit (Bioline) following the manufacturer's protocol. Detection of Wolbachia was carried out by real‐time PCR on a Rotor‐Gene
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Q machine (Qiagen) using strain‐specific primers and probes (Koga et al. 2009, Echeverria-Fonseca et al. 2015). Optimisation and conditions are detailed in Madhav et al. (2020b).
For quantification of Wolbachia single‐copy wsp genes were run in triplicate on a Rotor‐Gene Q machine using a SYBR® Green PCR Kit (Bioline). The relative density of Wolbachia was calculated using the 2–ΔΔCt method (Moreira et al. 2009). Methods are detailed in Madhav et al. (2020a)
2.3.2 Assay to visualise Wolbachia infection dynamics
Flourescence In Situ Hybridisation (FISH) assays wascarried out to visualise Wolbachia distribution in female BF post-adult microinjection using a method slightly modified from that of Koga et al. (2009). Briefly, for the whole-mount assay, 10 BF infected with wMel and wMelPop were collected six days post-injection and fixed in Carnoy’s solution overnight. Flies were sequentially washed the next day in 100%, 80% and 70% ethanol, and stored in 10% H2O2 in 100% ethanol for 30 days to quench the autofluorescence. Preserved flies were subsequently washed three times with 80% ethanol, 70% ethanol, and PBST× and pre-hybridised with hybridisation buffer (4× SSC, 0.2 g/ml dextran sulphate, 50% formamide, 250 μg/ml Poly A, 250 μg/ml salmon sperm DNA, 250 μg/ml tRNA, 100 mM DTT, 0.5× Denhardt’s solution) without probe two times for 15 min each. The insects were then incubated with hybridisation buffer and Wolbachia 16S rRNA probes overnight (Moreira 2009). The next morning, samples were washed three times with PBSTx, three times for 15 min each and finally incubated in PBSTx containing DAPI (10 mg/ml) for 30 min. Samples were then rewashed with PBSTx, covered with ProLong Diamond Antifade Mountant (Thermo Fisher Scientific, Brisbane, Australia) and photographed using a confocal microscope.
2.3.3 Wolbachia isolation from HIE-18 cells for microinjection
Wolbachia were isolated from the cell lines, according to Herbert & McGraw (2018). Briefly, wAlbB-, wMel- and wMelPop-infected cell lines were grown in 75 cm2 cell culture flasks for seven days using previously noted methods. Cells were pelleted on the eighth day by spinning at 2000×g and washed three times with SPG buffer (218 mM sucrose, 3.8 mM KH2PO4, 7.2 mM MK2HPO4, 4.9 mM L- glutamate, pH 7.5), sonicated on ice for two bursts of 10 s and cellular debris was removed by spinning at 1000×g for 10 min at 4 °C. The supernatant was passed through 50 μm and 2.7 μm acrodisc syringe filters (Eppendorf, Sydney, NSW, Australia) and centrifuged at 12,000×g to pellet Wolbachia. Finally, the pellet was suspended in 100 µl SPG buffer and used for microinjection.
2.4 Transinfection of BF with Wolbachia
2.4.1 Microinjection of eggs
The different stages of BF (eggs, pupae and adults) used for microinjection were sourced from our laboratory colony maintained at the EcoSciences Precinct and described earlier. To obtain eggs of similar age for microinjection, 7–10 day-old BF were held on moist filter paper in temporary cages for 20–30 min. Newly laid eggs (40–60 min-old) were arranged on double-sided sticky tape using a paintbrush and microinjected at the posterior pole of each egg with wAlbB (2 × 108 bacteria/ml) using a FemtoJet microinjector system (Eppendorf, Sydney, NSW, Australia). The microinjected eggs were then placed on tissue paper on the surface of manure pats to hatch.
After eclosion, larvae migrated into the moist manure where they fed until pupation. Pupae were separated from the manure by flotation in water on day 7 post-injection and incubated at room temperature. Flies that emerged from the puparium by day 10 were collected and separated by sex.
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Females that emerged from microinjected eggs were held singly with two uninfected males for mating in small cages made of transparent acrylic pipe (6 cm diameter × 15 cm height) closed with fly mesh and a membrane feeder at the top supplying cattle blood maintained at 26 °C. A 55 cm2 Petri dish containing moist filter paper was placed at the base of the cages for collection of eggs deposited by the flies. Females were allowed to oviposit and the eggs were collected until the death of the flies. Freshly deceased flies were collected and tested for the presence of Wolbachia using real-time PCR.
2.4.2 Microinjection of pupae
Approximately 3000–4000 eggs from colony-reared BF were incubated and the larva grown on manure to collect freshly pupated BF for microinjection (1–2 h-old). Pupae were aligned on double- sided sticky tape and injected in the third last segment at the posterior end close to germinal tissue using a FemtoJet microinjector system (Eppendorf). The microinjected pupae were then placed on moist Whatman filter paper and incubated at 27 °C until flies emerged. Freshly emerged flies were separated and placed in a cage with a maximum of five females and five males each. Eggs collected from each cage every day were tested for Wolbachia infection. Once infection was detected, female flies were separated into a separate single cage and eggs were collected for the G1 line until the flies died. Later, freshly deceased females were tested for the presence of Wolbachia using real-time PCR.
2.4.3 Microinjection of adults
Approximately 100–150 pupae from the BF colony at the EcoScience Precinct, Brisbane, Australia, were held separately from the main colony for collection of freshly emerged female flies (2–3 h-old) for injection. The female flies were collected within 3–4 h of eclosion from the pupae, anaesthetised 9 using CO2 for 30–40 s and then 2 µl of Wolbachia suspension (3 × 10 bacteria/ml) was injected into the metathorax of each fly using a handheld micro-manipulator (Burkard Scientific, London, UK) with hypodermic needles (0.24 × 33 mm). The microinjected flies (G0) were blood-fed and mated with male flies at the ratio of 1:1 in small cages as described above. On day three after injection, an artificial 100 g manure pat was placed onto sand at the base of each cage. Manure pats were removed every second day and the collected eggs were reared to adults following our standard laboratory protocols. Newly hatched G1 female flies were mated to potentially infected males, allowed to oviposit until death and the freshly deceased G1 flies then tested by real-time PCR for the presence of Wolbachia. Depending on the results of testing, the cycle was repeated.
2.5 Assessment of fitness effects
2.5.1 Survival
Two to three-hour-old post eclosion female BF were injected with Wolbachia (wAlbB, wMel and wMelPop) or SPG buffer (injected control) as described above and placed in triplicate cages containing ten flies each. Flies were cultured under laboratory conditions in small cages and mortality was noted every 12 h. Dead flies were later tested for Wolbachia infection individually using real-time PCR as described above. The survival assay for flies from microinjected pupae was carried as for adult-injected flies except that the number of flies in each cage was 20 (10 male and 10 female). Effects on the survival of BF injected with the different strains of Wolbachia and the SPG buffer control were analysed using an A log-rank (Mantel–Cox) test carried out using GraphPad Prism 8 software
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2.5.2 Adult emergence rate
Data from five independent pupae-microinjected batches were used to analyse the effect of Wolbachia on adult emergence. All three Wolbachia strains were injected in parallel to the buffer- injected controls. The number of injected pupae varied amongst batches between 77–205 for wMel, 98–145 for wAlbB and 82–148 for wMelPop. The emergence of adults was recorded each day and the ratio of total emerged to number of injected pupae was calculated to determine the final percentage of emergence. The effect of Wolbachia on pupal emergence was analyzed using one-way ANOVA followed by multiple comparison Tukey’s test using GraphPad Prism 8 software.
2.5.3 Egg production
The effect of Wolbachia on the number of eggs produced by females after pupal microinjection was assessed in triplicate with ten females per cage. Buffer-injected females were used as controls and number of eggs laid and females surviving were counted every 24 h to estimate eggs laid per day per female. If any females died during the assay, the number of eggs was adjusted to account for this. Dead females were later tested for the presence of Wolbachia using real-time PCR. One-way ANOVA followed by multiple comparison Tukey’s test was carried out using GraphPad Prism 8 software to test for difference in egg production.
3. Results and Discussion
3.1.1 Haematobia cell line establishment
The success of transinfection following microinjection is generally low, with subsequent loss of infection in newly infected hosts occurring commonly, particularly when transfer is between more distantly related host species (Anders et al. 2018). This is thought to be due to inability of the injected Wolbachia to adapt quickly to the new host environment. However, prior adaptation in cells more closely related to the target host species can increase the success rate of transinfection. For this reason The HIE-18 Haematobia cell line was established as previously described (Madhav2020a).
Figure 3. HIE 18 cell line. The cells are predominantly round, 5–10 μm, loosely adherent with ten mitotic chromosomes or tetraploid cells with 20 chromosomes. Arrow indicates a cell in metaphase (B);
3.1.2 Species confirmation using PCR and chromosomal analysis
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HIE‐18 cell identity was confirmed by amplifying the COI gene of ~ 920 bp with Haematobia‐specific primers. Both the HIE‐18 cells and positive control BF were amplified (Fig. 4A). HIE‐18 cells were comprised predominantly of round diploid cells (Fig. 3). Diploid cells had five pairs of chromosomes, two of which were sub‐metacentric and three metacentric (Figure 4B,C). There were no heteromorphic (sex) chromosomes. This line has now been successfully passaged 200 times in our laboratory and frozen down and successfully retrieved from liquid nitrogen storage on a number of occasions. Ten of the remaining 22 primary cultures developed into lines (passage seven or less) and are currently stored in liquid nitrogen.
Figure 4. PCR and chromosome profile confirmation of HIE‐18. (A) Approximate 920‐bp product amplified using mitochondrial‐encoded COI primer specific to Haematobia spp. HF, H. i. irritans HIE‐ 18 cell line; NTC, negative control; BF, H. i. exigua. (B) Metaphase chromosome spread in Giemsa‐ stained diploid HIE‐18 cells. (C) Plot shows a number of metaphase chromosomes in H. i. irritans cells.
3.1.3 Transinfection of cell line establishment and maintenance
3.1.3.1 Wolbachia replication in HIE‐18 cell lines
It took multiple introductions of Wolbachia into the HIE‐18 cell line flasks to establish persistent infections of wAlbB (two attempts), wMel (three attempts) and wMelPop (two attempts) and we saw a significant decrease in the density of Wolbachia within 48 h of infection: wAlbB (one‐way
ANOVA: F3,8 = 10.87, P = 0.0034), wMel (one‐way ANOVA: F3,8 = 27.30, P = 0.0001), and wMelPop
(one‐way ANOVA: F3,8 = 56.34, P < 0.0001; Fig. 5A–C). However, density subsequently increased and to date, HIE‐18 cells infected with wAlbB, wMel and wMelPop have been successfully maintained for 70, 60 and 50 passages, respectively, with stable infection and no significant change in Wolbachia density over these passages (Fig. 5D–F).
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Figure 5. Quantification of Wolbachia (wsp) relative to host (GAPDH) using real‐time PCR in HIE‐18 cell post infection. (A–C) Significant decreases in wAlbB, wMel and wMelPop were observed within 48 h post infection. *P < 0.02, **P < 0.003, ***P < 0.0003, ****P < 0.0001. (D) wAlbB dynamics in HIE‐18 cell line over 70 passages. (E) wMel dynamics in HIE‐18 cell line over 60 passages. (F) wMelPop strain dynamics in HIE‐18 cell line over 50 passages. There was no significant change in Wolbachia infection levels over succeeding passages P > 0.05 (D,E). Error bars indicate SEM from three culture flask replicates.
3.2 Microinjection
3.2.1 Microinjection of eggs
Embryonic microinjection is by far the most frequently used technique to develop Wolbachia transinfected insect lines, mainly because Wolbachia injected into the germ cells of the developing embryo provides a direct route for infection of the germ tissues in the early stage of differentiation (Hughes and Rasgon 2014). Of a total of 2036 eggs microinjected with the wAlbB strain, only 10 developed through to adult flies (six females and four males) and no infection was detected in any of the adults. Microinjecting BF was found to be particularly difficult because of the tough chorion surrounding the egg that is formed during embryogenesis. This proved to be a significant barrier to successful microinjection, causing rapid blunting and frequent breaking of the needles, frequent visible damage to the eggs and based on the hatch rate, a high level of damage to the embryos.
We measured significant reductions in embryo survival and hatching from the effects of injection (Fisher’s exact test: P < 0.0001) (Fig. 6a) and identified that older eggs (40–60 min) had a better injection survival rate, 21.96% compared to 3.4% for younger eggs (10–30 min) (Tukey’s multiple comparison test: P = 0.010) (Fig. 6b). A number of variations of the technique were tested to improve the survival rate of eggs post-microinjection. These included dechorionation of the eggs with 2.5% sodium hypochlorite for 30 s to soften the chorion, partial desiccation to reduce hydrostatic pressure in the eggs and increase space for the retention of larger volumes of injectate
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and the use of halocarbon oil (2:1 mix of halocarbon 700 and 27) to prevent desiccation of the eggs. None of these treatments markedly improved egg/larvae survival post-microinjection (2.33%) and they also appeared to reduce egg survival in uninjected eggs (16.33%) (one-way ANOVA: F(2,
6)= 181.6, P < 0.0001) (Fig. 6c). Another disadvantage of this technique is the inability to determine the sex of an embryo prior to injection which means that approximately half of the injected flies will be males that do not transmit Wolbachia to the next-generation (Hughes and Rasgon 2014) Similar difficulties with microinjection were experienced when attempting to use microinjection for gene transfection in closely related H. i. irritans eggs. In this instance, the researchers opted to use electroporation, which is unsuitable for the introduction of bacteria (Xu et al. 2016)
Figure 6. Challenges with BF embryonic microinjection. a. Sham Embryonic microinjection (without Wolbachia) had a detrimental effect on embryo hatching. b. 40–60 min-old embryos survived injection better than 10–30 min-old embryos. c. Eggs were dechorionated by treating with 2.5% sodium hypochlorite for 30 s and covered with 2:1 mix of halocarbon oil 700 and 27 to prevent desiccation. Eggs were sensitive to treatment and survival decreased further with injection. Error bars are SEM. Analysis was by Fisher’s exact test in (a) and Tukey’s multiple comparison test in (b) and (c). ****P < 0.0001)
Table 1: Egg microinjection survival and infection rate
Wolbachia No. eggs No. of adult Infection strain injected No of pupae flies emerged percentage wAlbB 12 batches 2123 10 7 Adult: 0/7 Adult: 0/17 wMel 5 batches 1760 27 17 Egg: 0/30 wMelPop 7 Batches 3421 58 41 Adult: 0/41
3.2.2 Microinjection of adults
Although embryonic microinjection has been the primary method used to transinfect insects, adult microinjection can be advantageous in that females can be selected, increasing the efficiency of injecting (Hughes and Rasgon 2014). Further, adult microinjection can be performed using a simple syringe and small-bore needles delivering higher volumes of Wolbachia to overcome the host immunological response (Hughes and Rasgon 2014).
Table 2. Adult microinjection: Wolbachia infection by generation (G1=injected)
Number Generation - Infected/ total tested Strain Injected G1 G2 G3 G4
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Adult: Adult: 5/89 378 Adult NA wAlbB 118/126 (5.6%) (19 batches) Eggs: 0/50 (95.93%) (1 batch) Adult: Adult: Pupae 441 27/119 Adult: 0/25 wMel 117/123 12/24 (17 batches) (22.68%) Eggs: 0/100 (95.12%) (4 batches) (50%) Adult: Adult: 10/91 Adult: 2/60 417 wMelpop 103/106 (10.98 %) (3.3%) (15 batches) (96.26%) (4 batches) Eggs NA
3.2.2.1 Growth of Wolbachia in injected females
The growth kinetics of Wolbachia were studied in injected female flies by quantifying Wolbachia on days 3–11 compared to day 0 (day of injection). Overall, the pattern showed an initial significant decrease in Wolbachia density to approximately day 5 followed by subsequent growth and increase in bacterial titre to day 11 in all three strains (Kruskal–Wallis H-test: χ2= 31.18, df = 5, P < 0.0001) (Fig. 7a–c). It is notable that although all three strains were injected with the same amount of Wolbachia, but the level of wMelPop, as measured by RT-PCR was 50% higher at the end of 11 days a than wAlbB. This was not unexpected as wMelPop is known to be much more aggressive and to grow much faster in other insects as well (Ritchie et al. 2015).
Figure 7: Wolbachia dynamics post-adult microinjection of female BF assessed using real-time PCR. Mean Wolbachia density is expressed relative to the host genome. Kruskal–Wallis test and Dunn’s multiple comparison test were used to compare titres at day 0. All error bars are SEM. Bars with different letters in each graph are significantly different (p< 0.05).
3.2.2.2 Distribution of Wolbachia infection in injected female BF
Difference in the quantitative dynamics of Wolbachia growth after injection, particularly the spatial dynamics of infection in injected flies, required a better understanding of tissue tropism. Hence, fluorescence in situ hybridisation (FISH) was carried out on whole mounted BF and dissected ovaries to visualise the localisation of wMel and wMelPop Wolbachia six days after injection (Fig. 8). When this was done, Wolbachia infection was shown to be widely distributed and clearly growing in somatic tissues, including the thoracic muscle, head, abdominal area, proboscis and legs (Fig. 8). However it was not found in the germline tissues which is required for transmission of Wolbachia to the next generation. The PCR results for Wolbachia growth in flies (Figs. 8, 9) suggest that the use of FISH at 6 days post-injection was too early to determine the final distribution of Wolbachia. Hence, we studied tissue invasion and the detailed distribution of Wolbachia in adult flies by real-time PCR after dissecting out the thoracic muscle, midgut, fat bodies, ovary and head at nine days post-adult
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injection (Fig. 4a–c). Wolbachia were found to be replicating in all somatic tissues with wAlbB having an infection percentage of 33–83% (n = 6) and wMel and wMelPop between 66–100% (n = 6). No infection was found in germline tissues. However, on a few occasions first-generation flies from adult injection with wAlbB, wMel,and wMelPop were found positive with infection percentages of 5%, 22% and 10%, respectively, indicating transmission via the germline tissues in these instances (see Table 2).
Figure 8. Fluorescence in situ hybridisation images showing localisation of Wolbachia six days post- adult injection. Wolbachia is distributed throughout the fly (blue: host, pink/red: Wolbachia): a) wMel in head and thorax. b) wMelPop in the abdominal region. c).wMelPop in the head, mouthparts, thorax and leg. d) Control no bound fluorescence. Abbreviations: T, thorax; H, head; A, abdomen; M, mouthparts; L, leg
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B.AHE.0242 – Wolbachia control of buffalo fly
Figure 9: Wolbachia tropism post-adult microinjection of female BF assessed using real-time PCR. Wolbachia tropism in female (n = 6) nine days post-adult injection (a–c). None of the Wolbachia strains was found in the ovaries.
3.2.2.3 Survival of Wolbachia infected females:
The results of the survival assays showed that by day seven less than 20% of the wMelPop and less than 50% of wMel and wAlbB injected flies were alive (Fig. 10) whereas more than 90% of buffer- injected flies were alive. Both wMelPop (log-rank statistic= 16.92, P < 0.0001) and wMel (log-rank statistic= 11.96, P = 0.0005) significantly reduced longevity of female BF. However, there was no significant effect of the wAlbB strain in comparison to the control injected flies (log-rank statistic= 0.25, P = 0.62).
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Figure 10: Survival of female BF post-adult injection with Wolbachia. Triplicate cages of adult flies each containing ten females were maintained under lab culturing conditions. The number of dead flies were recorded until all died. A significant reduction in survival was observed in wMel (P < 0.0005) and wMelPop (P < 0.0001) injected flies by Log-rank (Mantel–Cox) tests
Our results with adult injection of Wolbachia were promising. Despite that injections in first few batches were made mainly with Wolbachia grown in D. melanogaster cells (wAlbB, wMel and wMelPop strain), not previously adapted in Haematobia cells, infection rates and persistence in the injected flies were high (generally > 90%). In some batches, transmission to the next-generation was confirmed.
3.3 Microinjection of pupae
As oviposition by BF may begin as early as three days after eclosion from the pupae and continue until death, confirmation that Wolbachia was actually growing in the flies and knowledge of Wolbachia distribution and dynamics in injected females was important to identify the optimal time to collect eggs for the establishment of infected colonies. The optimal time was identified as 11-15 days of age at which time growth of Wolbachia was maximised, but age-related mortality was still low.
Table 3: Pupal microinjection: Wolbachia infection by generation (G1=injected)
Total Generation - Infected/ total tested Strain injected G1 G2 G3 G4 Adult : 895 119/132 Adult: 0/20 Adults: 0/15 Adult: 0/13 wAlbB (10 batches) (90.15%) (0 %) (0%) (0%) Egg: 10/80 (2 batches) Egg: 0/50 Eggs: 0/140 Egg: 0/60 Adult: 1820 112/123 Adult: 2/9 Adult: 0/8 Adult: 0/8 wMel (18 batches) (91.10 %) (22%) (0%) (0%) Egg: 30/400 Egg: 10/120 (3 batches) (1 batch) Egg: 0/80 Egg: 0/90 Adult: 2141 148/162 Adult: 0/23 Adult: 0/14 Adult: 0/10 wMelpop (20 batches) (91.35 %) (0%) (0%) (0%) Egg= 55/520 Egg: 10/150 6 batches) (1 batch) Egg: 0/50 Egg: 0/40
3.3.1 Wolbachia growth and tissue distribution in pupal –infected BF
Wolbachia density significantly decreased to day five, likely due to host immune response (Madhav et al. 2020) but recovered by day eleven after injection. A similar result was obtained when wMelPop and wAlbB were injected into Anopheles gambiae adult mosquitoes (Hughes et al. 2011) The initial host immune response was anticipated as the densities of wAlbB, wMel and wMelPop Wolbachia in the Haematobia cells were also observed to decrease initially, possibly due to an innate immune response mediated by the Imd pathway (Madhav et al. 2020). Real-time PCR analysis of dissected tissues nine days after injection showed Wolbachia to be present in all the vital somatic
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tissues (head, thoracic muscle, midgut and fat body), but not the ovarian tissues, suggesting that Wolbachia might need extra time to infect the ovaries. However, injection with wAlbB, wMel and wMelPop Wolbachia caused > 40% death in flies by day seven post-injection, reducing the time window for collecting infected eggs. Therefore, we hypothesised that microinjecting 1–2 h-old pupae would give more time than with adult microinjection for Wolbachia to multiply, spread and establish in the ovaries. Pupal injection has previously been conducted with Trichogramma wasps and resulted in successful ovarian infections and persistence of Wolbachia in the wasp colony for 26 generations [60].
In this case, wMel and wMelPop overcame host immune responses and established in both somatic and germline tissues. Further, in two instances, next-generation (G1) BF from wAlbB and wMel injected pupae were positive for Wolbachia, indicating next-generation transmission as a result of pupal injection. The main disadvantages of pupal injection in comparison with adult injection were limitation on the volume of Wolbachia that could be injected and inability to distinguish female from male pupae for injection.
A similar quantitative assay to that used for injected adult BF was carried out to track the dynamics and tropisms of the three Wolbachia strains post-pupal injection. The extra time in the pupal phase resulted in 66–100% infection in the somatic tissue with wAlbB and wMel (n = 6) and 83–100% with wMelPop (n = 6) 13 days post-pupal injection (Fig. 11a–c). Furthermore, in 16% of cases the ovaries of females injected with wMel and wMelPop Wolbachia were found to be infected. Also, two first- generation flies from wMel-injected pupae and four eggs from wAlbB-injected pupae were found positive for Wolbachia infection (Table 3). Analysis of Wolbachia dynamics showed approximately the same pattern as for adult injection, where density initially decreased in the first seven days, then significantly recovered by day 9 in wMel (Kruskal–Wallis H-test: χ2= 29.61, df = 5, P < 0.0001) and day 13 in wMelPop and wAlbB post-pupal injection (Kruskal–Wallis H-test: χ2= 32.12, df = 5, P < 0.0001) (Fig. 611 d–f).
Figure 11. Wolbachia tropism and dynamics post-pupal microinjection of female BF assessed using real-time PCR. a–c Wolbachia tropism in female BF (n = 6) 13 days post-pupal injection. Ovary
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infection was detected in wMel and wMelPop injected flies. d–f Wolbachia dynamics measured over 15 days post-injection. Here, mean Wolbachia density is expressed relative to the host genome. Kruskal–Wallis and Dunn’s multiple comparison tests were used to compare titres to those at day 0. Bars with different letters are significantly different (P < 0.05). Scale on the Y-axis for wMelPop (f) is different to that for the other two strains (d, e) indicating faster growth rate with wMelPop
3.3.1 Effect of pupal injection with Wolbachia on survival of emergent BF
A significant decrease in the longevity of BF post-pupal injection was found in both sexes of wMelPop-injected BF (male: log-rank statistic= 20.25, P < 0.0001, female: log-rank statistic= 29.04, P < 0.0001), but the effect was not significant with the two other strains [wAlbB: male (log-rank statistic= 2.267, P = 0.132), female (log-rank statistic= 3.275, P = 0.071); wMel: male (log-rank statistic= 3.027, P = 0.1545), female (log-rank statistic= 3.467, P = 0.063)] (Fig. 7).
Figure 12: Survival of BF post-pupal injection with Wolbachia. Triplicate cages of flies eclosed from pupae on the same day (10 males and 10 females per cage) were maintained in laboratory culturing conditions. Mortality was recorded daily until all flies were dead. Log-rank (Mantel–Cox) showed a significant reduction in the male wMelPop (P < 0.0001) and female wMelPop (P < 0.0001) injected flies
The wMelPop strain is a virulent type of Wolbachia, and its over replication in somatic tissues and brain cells, known in other infected insects (Pietri et al. 2016, Min and Benzer 1997)), may have been the reason for the early death of BF. Further, in the studies of Wolbachia kinetics we found a higher density of wMelPop than with the other two strains following both adult and pupal injection. Reduction in the longevity of infected Ae. aegypti mosquitoes caused by infection with wMelPop, decreasing the potential extrinsic incubation time for the dengue virus, was one of the characteristics that led to the hypothesis that wMelPop infection would reduce dengue spread (McMenniman 2009) Infection with wMelPop could also markedly reduce BF life span and their ability to transmit Stephanofilaria sp. nematodes. These nematodes have been implicated in the development of BF lesions, a significant production and welfare issue in north-Australian cattle (Shaw and Sutherland 2006). Stephanofilaria has an extrinsic incubation period of up to 3 weeks in Haematobia spp. (Hibler 1966) and the life-shortening effects of Wolbachia shown in our study could markedly reduce the vector competency of infected flies. There is also the possibility the Wolbachia infection could more directly compromise the vector competency of BF for Stephanofilaria sp., as has been seen in the case another filarial nematode, Brugia pahangi transmitted by mosquitoes and in the case transmission of the dengue virus by Ae. aegypti [Kambris et al. 2009, Walker et al. 2011].
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3.3.2 Effect of pupal injection with Wolbachia on adult emergence
Infection of the somatic tissues by Wolbachia can have consequences on physiological processes. Non-injected control flies emerged from pupae after 3–7 days, whereas mock-injected control flies emerged from 5 to 7 days, wAlbB after 6–7 days and wMel- and wMelPop-injected flies at 5–7 days post-injection (Fig. 13a). It is important to note that emergence in wMel- and wMelPop-injected flies was less than 2% on day 5. Overall, there was significant decrease in the percent emergence of wMel-injected (30.01 ± 3.91) (Tukey’s multiple comparison test, P = 0.0030) and wMelPop-injected flies (27.98 ± 3.92) (Tukey’s multiple comparison test, P = 0.0011) compared to the control injected flies (46.95 ± 4.15), but no significant difference was observed with the wAlbB-injected flies (Tukey’s multiple comparison test: P = 0.77) (Fig. 13b). Nearly 5% of the flies that emerged from the wMelPop-injected pupae were too weak to completely eclose from the pupal case and had deformed wings (Fig. 13 c, d).
Figure 13. Fitness effects on buffalo fly post-pupal injection with Wolbachia. a Wolbachia delayed adult emergence. b A significant decrease in adult emergence was observed in wMel (P = 0.0030) and wMelPop (P = 0.0011) injected pupae when analysed using Tukey’s multiple comparison test. Nearly 5% of wMelPop flies either failed to completely eclose from the pupal case or had deformed wings
3.3.4 Effect of pupal injection with Wolbachia on egg production by BF
Difference between infected females and non-infected females in egg production was also analysed following pupal injection with the three different strains of Wolbachia. Over 14 days there was a significant reduction in the total number of eggs laid by females infected with wAlbB (P = 0.012), wMel (P = 0.0052) and wMelPop (P = 0.0051) in comparison with the mock-injected flies (Fig. 14).
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Figure 14. Fecundity of BF post-pupal injection with Wolbachia. Flies started laying eggs from day 3 post-emergence and continued until day 16. Eggs laid from triplicate cages each having ten females was recorded every day for (a) wAlbB (b) wMel and (c) wMelPop. (d) A significant reduction in the total number of eggs laid per female over 13 days was found in flies infected with wAlbB (P = 0.0123), wMel (P = 0.0052) and wMelPop (P = 0.0051) (Tukey’s multiple comparison test)
Fecundity of insects has a significant influence on the population dynamics of insect populations and when Wolbachia-infected males are released CI, vertical transmission, and a relatively higher fertile egg production by infected females in comparison with uninfected flies increases the chances of successful establishment of Wolbachia in a new host population (Weeks et al. 2007) Wolbachia have been found to enhance and reduce egg production depending upon both the strain of the Wolbachia and the host (Rigaud et al. 1999, Fleury et al 2000, Grenier et al. 2002, Dobson et al. 2004, Weeks et al. 2007, Sarakatsanou 2011, Joshi et al 2014). We found that wAlbB, wMel and wMelPop significantly reduced total egg production in pupal injected flies (Fig. 14d). In addition, Wolbachia infection caused delayed and decreased adult emergence of BF post-pupal injection. Wolbachia being an endosymbiont lacks nutritional biosynthetic pathways and depends on its host for wide range of nutrition (Foster et al. 2005, Caragata et al. 2014). Hence, the fitness costs observed in injected BF could be the result of competition between high density of Wolbachia and BF for nutritional resources such as amino acids and lipids (Caragata et al. 2014]. Another possibility could be that as Wolbachia was found in all of the critical tissues involved in the endocrine cascades for egg production and maturation in insects (midgut, neuron, fat bodies and ovary), it interfered with egg production by this means (Negri et al. 2012). In addition, delayed larval development associated with wMelPop infection has been documented in mosquitoes on a number of occasions (McMeniman and O’Neill 2010). If these deleterious effects are a consistent feature of Wolbachia
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infection in BF, they could have a significant impact in altering population dynamics or even crashing BF populations [McMeniman and O’Neill 2010, Ritchie et al 2015]. For instance, female BF lay eggs in fresh cattle manure pats, where eggs take approximately seven days to develop into pupae depending upon the temperature and moisture content of the pat (Cook and Spain 1982). Prolonged larval development and time to eclosion of Wolbachia-infected BF, together with adult life span reduction might decrease overwintering and survival of BF, particularly during periods of unfavourable fly conditions and at the edge of the BF range.
4 Conclusion The development of this approach requires a lot of steps to be successfully completed. Firstly, nearly all previous research with BF has been conducted using colonies of flies reared on live cattle (Stegeman et al. 1996). Although we contemplated the possibility of conducting this project using flies caged on the backs of cattle using a rearing system such as this, with the myriad tests and intermating of flies required and the need to maintain colonies long term this was not really feasible. In addition, animal ethics considerations and approvals come into play when cattle are used for this purpose. Therefore the first tool that was needed was a completely in vitro colony of BF, rather than a colony that used live cattle to rear BF as previously used. This colony, the only one in Australia and indeed the world, has now been maintained through more than 180 generations in our laboratory. This was an important step and is critical tool for this, or like projects to proceed.
The second step was to develop a BF cell line, to adapt Wolbachia originally isolated from Drosophila melanogaster (wMel, wMelPop) and Aedes albopictus (wAlbB) to the Haematobia context to increase the likelihood of successful transinfection of BF (McMeniman et al 2009). It should be noted, however, that although we had tested Australian populations of BF and determined that Wolbachia was not currently present (Zhang et al. 2009) we knew that Wolbachia was present in very closely related HF and thus we expected BF to be a similarly competent host. We had previously established a BF cell line during the 2010 floods and despite multiple attempts in the current project have had difficulty re-establishing this line. A planned collaboration with Profs Tim Kurtti and Uli Munderloh at the University of Minnesota to this end as part of this project was held up by delays in AQIS approvals to import flies from our colony to the biosecurity facility at the University of Minnesota. However, while waiting for the approval we established the current H.i. irritans (HIE-18) line (as far as we know currently the only one in the world) (Madhav et al. 2020a). Given the extremely close relatedness of the two sub species (Iwasa and Ishigura 2010) we decided to proceed using this line. We have now successfully transinfected this line with the three strains of Wolbachia and the wAlbB wMel and wMelPop infected lines have now been successfully grown through 125, 112 and 98 passages respectively. It should be noted that the loss of infection in newly infected hosts is common, particularly in more distantly related host species (McMeniman 2008, 2009) and many of the injections described in this study were conducted with Wolbachia early in the adaptation process. The further adaptation of Wolbachia in the Haematobia cells as a result of further passaging should also help to increase the likelihood of sustaining infections developing from future injections.
The main approach used for transinfection of insects with Wolbachia to date has been embryonic (egg) microinjection whereby 0.5 m gauge needles are inserted through the egg chorion and Wolbachia are introduced directly into the egg cytoplasm, maximising the likelihood of transmission to succeeding generations. However, the tough outer covering of the eggs presented significant difficulties for BF microinjection causing rapid blunting and frequent breakage of microinjection needles and significant trauma to the eggs. This resulted in very low hatch rates with no successful transmission to the next generation detected. Of a total of 7304 eggs injected only 95 pupae were collected, 65 of those developed to adult flies and none had Wolbachia detected. Similar difficulties
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were also encountered by by Xu et al. (2016) when attempting to microinject genetic material into HF eggs.
We therefore tested adult microinjection whereby recently emerged BF were injected. Injection was followed by an initial reduction in Wolbachia numbers, presumably as a result of host immune response. However this was followed by recovery in Wolbachia numbers and subsequent growth which resulted in widespread somatic infection, confirming BF as a competent host the three Wolbachia strains used. Although we did not detect ovarian infection in the FISH studies, only limited numbers of insects were tested by this method and the detection of Wolbachia in G2, G3 and in one instance G4 generations in the adult injected stocks indicated that ovarian infections had occurred on a number of occasions. In addition, it took some time for the injected Wolbachia to overcome the insect immune response so that by the time Wolbachia had build up to levels where ovarian infection was likely to occur the insect was nearing the end of its life, providing a relatively short window for the production of infected eggs.
Therefore, to prolong the time for the Wolbachia to build up, and to increase the opportunity for the production of infected eggs, we also tested pupal microinjection. This was shown to provide earlier build up and higher levels of Wolbachia infection, and PCR testing indicated ovarian infections in many of the hatching flies. Subsequent microinjection has also resulted in the detection of infection in G2 eggs, also indicative of ovarian infection.
The results with the adult and pupal microinjection have been extremely encouraging although we are yet to achieve sustainably Wolbachia-infected strain of BF. On reflection we perhaps spent too much time in this project trying to adapt the embryonic microinjection technique and more time refining the adult and pupal injection techniques would have been a better option. Achieving successful transinfections is usually a tedious process and that we have not yet achieved a sustainably infected strain is perhaps unsurprising given the relatively small number of pupal and adult injections that we have conducted as a result. As an example, microinjection of 10,000 A. aegypti eggs was required for the establishment of two Wolbachia-infected mosquito lines that formed the basis of the world recognised denque and zika virus control programs (McMeniman et al. 2009, Ritchie et al. 2014). However, despite that we have conducted relatively few attempts using pupal and adult microinjection, we have now achieved ovarian infections and transmission across generations on a number of occasions. Using these methods and together with better adapted strains and the rapid advances in Wolbachia technology currently being made in other labs we believe that there is a high probability that further work in this area will yield a successful outcome.
It should be stressed that this is a high risk – high reward approach. Most current animal pest and parasite control programs rely on direct animal treatments. These measures are expensive, often labour intensive and when chemical insecticides are used, risks the development of resistance, occupational health and safety exposure, residues in produce and environmental contamination. The approach suggested here, directly targeting pest populations rather than using on-farm animal treatments would provide significant labour savings and would enhance to reputation of Australian produce as clean, safe and ethically produced. Further, BF are continuing to extend their range southwards and the impacts of BF are expected to increase under the effects of climate change. A Wolbachia-based approach offers a novel, convenient and practicable approach to arresting the range expansion of BF where chemical treatments and regulated cattle movements have previously proven unsuccessful (Williams et al. 1985) and the potential for suppression or local elimination of BF or BF lesions in endemic areas. The very favourable potential cost benefit provided by area wide approaches is well illustrated by the sterile male approach that eradicated screwworm from North and Central America where the direct benefits achieved each year from the programme are greater
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than the total cost of the programme over the fifty years of its operation. A Wolbachia-based approach offers the added advantage that as Wolbachia can drive itself through populations from relatively low infection levels, very much lower release ratios are required than with other genetic approaches.
4.1 Key findings
• We have developed a number of critical resources to enable development of a Wolbachia based, area wide approach to BF control: ▪ A completely in- vitro buffalo fly colony now maintained in the laboratory for 191 generations and critical for this work ▪ A Haematobia cell line for adaption Wolbachia to the BF cellular context. Cells from the strain have been stored in liquid nitrogen and subsamples have been re-established on a number of occasions, demonstrating viability ▪ Three Haematobia-adapted strains of Wolbachia for microinjection into BF • wAlbB (grown though 125 passages) • wMel (128 passages) • wMelPop (98 passages) These strains have now been grown through more than 100 passages in Haematobia cells and should be well adapted for transinfection into BF • We have shown that BF is a competent host for Wolbachia and that all three Wolbachia strains tested successfully infect and grow in injected BF.
• We have developed pupal and adult microinjection techniques that can produce ovarian infections of Wolbachia that are transmitted across generations and provide a more tenable approach to develop sustainably infected strains of BF than embryonic microinjection.
• Pupal injection produced higher and earlier build-up of Wolbachia infection levels than adult injection, and showed ovarian infections in a number of instances
• Infection with Wolbachia caused reduction in lifespan, reduced fecundity, reduced and delayed pupal emergence
• In view of these results, we are now well placed to establish Wolbachia-transinfected BF lines suitable for use in future biocontrol programs for BF population suppression.
4.2 Benefits to industry
Control of BF is increasingly compromised by the development of resistance to control products and there are rapidly growing markets for meat and milk produced in low- chemical systems. The move towards a greater proportion of Bos taurus genes in northern cattle herds to raise beef quality will increase susceptibility to BF. In addition, BF has been spreading steadily southward and global warming is predicted to markedly increase the range of BF. CLIMEX modelling suggests that sustaining resident populations could establish through most of the coastal belt of NSW, in moister areas of SA and in southern WA by as soon as 2030, with profound effects on the non-tolerant B. taurus breeds on which beef and dairy production is based in these areas.
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Most current BF control programs rely on direct animal treatments. These measures are expensive, often labour intensive and when chemical insecticides are used, risks the development of resistance, occupational health and safety exposure, residues in produce and environmental contamination. The approach suggested here, directly targeting pest populations rather than using individual animal treatments would provide significant labour savings on-farm and would enhance to reputation of Australian produce as clean, safe and ethically produced. Further, BF are continuing to extend their range southwards and the impacts of BF are expected to increase under the effects of climate change. A Wolbachia-based approach offers a novel, convenient and practicable approach to arresting the range expansion of BF where chemical treatments and regulated cattle movements have previously proven unsuccessful, as well as a pathway to area-wide suppression or elimination of BF populations.
5 Future research and recommendations
• We have shown that adult and pupal microinjection are much more effective and practicable methods than embryonic microinjection for the transinfection of BF. We have also shown that BF is a competent host for Wolbachia and have generated ovarian infections and transmission to succeeding generations on a number of occasions in this project, but because we spent so much time on trying to perfect the embryonic technique, we have not had time to ‘tweak’ the pupal/adult techniques to develop sustainable infections. Advice is that this should be feasible and with the knowledge gained in this project there is high likelihood of successful establishment of sustaining Wolbachia-infected BF strains. There would be benefits in partnering with a specialist Wolbachia research lab towards this end.
• Wolbachia based approaches could be used in three main ways: ▪ The use of direct fitness effects and fertility penalty to reduce population build up and survival, particularly in overwintering foci at the edge of the BF range, towards potential local eradication of buffalo flies and the prevention of southerly spread ▪ Reduction in the transmission of Stephanofilaria, towards reducing the occurrence of buffalo fly lesions ▪ Incompatible Insect Technique that could be used to eradicate foci for summer spread of BF and foci of BF invasion into southern areas
• A buffalo fly population model should be developed to design and evaluate optimal Wolbachia- based control approaches
• The potential to use the wirr strain of Wolbachia, which is widely spread in closely related HF (Haematobia irritans irritans) in control programs for BF should also be investigated
6 References
Alam, U., J. Medlock, C. Brelsfoard, R. Pais, C. Lohs et al., 2011 Wolbachia symbiont infections induce strong cytoplasmic incompatibility in the tsetse fly Glossina morsitans. PLOS pathog 7: e1002415. Anders, K. L., C. Indriani, R. A. Ahmad, W. Tantowijoyo, E. Arguni et al., 2018 The AWED trial (Applying Wolbachia to Eliminate Dengue) to assess the efficacy of Wolbachia-infected mosquito deployments to reduce dengue incidence in Yogyakarta, Indonesia: study protocol for a cluster randomised controlled trial. Trials 19: 1-16. Andrews, E. S., P. R. Crain, Y. Fu, D. K. Howe and S. L. Dobson, 2012 Reactive oxygen species
Page 37 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
production and Brugia pahangi survivorship in Aedes polynesiensis with artificial Wolbachia infection types. PLoS pathogens 8: e1003075. Animal Health Australia, 2017. Screwworm fly surveillance and preparedness program. Animal Health Australia, Canberra Australia. https://www.animalhealthaustralia.com.au/what-we-do/disease- surveillance/screw-worm-fly. Anstead, C. A., P. K. Korhonen, N. D. Young, R. S. Hall, A. R. Jex et al., 2015 Lucilia cuprina genome unlocks parasitic fly biology to underpin future interventions. Nature communications 6: 1-11. Bean, K., G. Seifert, A. Macqueen and B. Doube, 1987 Effect of insecticide treatment for control of buffalo fly on weight gains of steers in coastal central Queensland. Australian Journal of Experimental Agriculture 27: 329-334. Bian, G., G. Zhou, P. Lu and Z. Xi, 2013 Replacing a native Wolbachia with a novel strain results in an increase in endosymbiont load and resistance to dengue virus in a mosquito vector. PLoS Negl Trop Dis 7: e2250. Blagrove, M. S., C. Arias-Goeta, A.-B. Failloux and S. P. Sinkins, 2012 Wolbachia strain wMel induces cytoplasmic incompatibility and blocks dengue transmission in Aedes albopictus. Proceedings of the National Academy of Sciences 109: 255-260. Bordenstein, S. R., and S. R. Bordenstein, 2017 Phage-mediated manipulation of wolbachia, pp. Google Patents. Bourtzis, K., R. S. Lees, J. Hendrichs and M. J. Vreysen, 2016 More than one rabbit out of the hat: Radiation, transgenic and symbiont-based approaches for sustainable management of mosquito and tsetse fly populations. Acta tropica 157: 115-130. Brelsfoard, C. L., W. St Clair and S. L. Dobson, 2009 Integration of irradiation with cytoplasmic incompatibility to facilitate a lymphatic filariasis vector elimination approach. Parasites & vectors 2: 38. Calvitti, M., R. Moretti, E. Lampazzi, R. Bellini and S. L. Dobson, 2014 Characterization of a new Aedes albopictus (Diptera: Culicidae)-Wolbachia pipientis (Rickettsiales: Rickettsiaceae) symbiotic association generated by artificial transfer of the w Pip strain from Culex pipiens (Diptera: Culicidae). Journal of medical entomology 47: 179-187. Caputo, B., R. Moretti, M. Manica, P. Serini, E. Lampazzi et al., 2020 A bacterium against the tiger: preliminary evidence of fertility reduction after release of Aedes albopictus males with manipulated Wolbachia infection in an Italian urban area. Pest Management Science 76: 1324-1332. Caragata, E. P., and L. A. Moreira, 2017 Using an endosymbiont to control mosquito-transmitted disease, pp. 123-142 in Arthropod Vector: Controller of Disease Transmission, Volume 1. Elsevier. Caragata, E. P., E. Rancès, L. M. Hedges, A. W. Gofton, K. N. Johnson et al., 2013 Dietary cholesterol modulates pathogen blocking by Wolbachia. PLoS Pathog 9: e1003459. Caragata, E. P., E. Rancès, S. L. O’Neill and E. A. McGraw, 2014 Competition for amino acids between Wolbachia and the mosquito host, Aedes aegypti. Microbial ecology 67: 205-218. Champer, J., A. Buchman and O. S. Akbari, 2016 Cheating evolution: engineering gene drives to manipulate the fate of wild populations. Nature Reviews Genetics 17: 146. Chen, L., C. Zhu and D. Zhang, 2013 Naturally occurring incompatibilities between different Culex pipiens pallens populations as the basis of potential mosquito control measures. PLoS Negl Trop Dis 7: e2030. Cook, I., and A. Spain, 1982 The Effects of Temperature and Moisture on Survival of the Immature Stages of the Buffalo Fly, Haematobia Irritans Exigua De Meijere (Diptera: Muscidae). Australian
Page 38 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
Journal of Zoology 30: 923-930. Curtis, C., and S. Sinkins, 1998 Wolbachia as a possible means of driving genes into populations. Parasitology Cambridge- 116: S111-S115. Department of Agriculture and Water Resources Australia, 2017, State and Territory notifiable animal diseases list. Department of Agriculture and Water Resources, Canberra, Australia. http://www.agriculture.gov.au/pests-diseases-weeds/animal/state-notifiable de Azambuja Garcia, G., G. Sylvestre, R. Aguiar, G. B. da Costa, A. J. Martins et al., 2019 Matching the genetics of released and local Aedes aegypti populations is critical to assure Wolbachia invasion. PLoS neglected tropical diseases 13: e0007023. De Barro, P. J., B. Murphy, C. C. Jansen and J. Murray, 2011 The proposed release of the yellow fever mosquito, Aedes aegypti containing a naturally occurring strain of Wolbachia pipientis, a question of regulatory responsibility. Journal für Verbraucherschutz und Lebensmittelsicherheit 6: 33-40. Dobson, S., W. Rattanadechakul and E. Marsland, 2004 Fitness advantage and cytoplasmic incompatibility in Wolbachia single-and superinfected Aedes albopictus. Heredity 93: 135-142. Dowell, F., A. Parker, M. Benedict, A. Robinson, A. Broce et al., 2005 Sex separation of tsetse fly pupae using near-infrared spectroscopy. Dyck, V. A., J. Hendrichs and A. S. Robinson, 2006 Sterile insect technique: principles and practice in area-wide integrated pest management. Springer. Echeverría-Fonseca, G., P. A. Mera-Ruiz, J. Carrillo-Toro and R. Rodriguez-Hidalgo, 2015 A new DNA extraction protocol for screwworm fly Cochliomyia species (Diptera: Calliphoridae). Frontiers in Environmental Science 2: 68. Fleury, F., F. Vavre, N. Ris, P. Fouillet and M. Bouletreau, 2000 Physiological cost induced by the maternally-transmitted endosymbiont Wolbachia in the Drosophila parasitoid Leptopilina heterotoma. Parasitology 121: 493-500. Floate, K. D., G. K. Kyei-Poku and P. C. Coghlin, 2006 Overview and relevance of Wolbachia bacteria in biocontrol research. Biocontrol Science and Technology 16: 767-788. Foster, G., W. Vogt and T. Woodburn, 1985 Genetic analysis of field trials of sex-linked translocation strains for genetic control of the Australian sheep blowfly Lucilia cuprina (Wiedemann). Australian journal of biological sciences 38: 275-294. Foster, G., W. Vogt, T. Woodburn and P. Smith, 1988 Computer simulation of genetic control. Comparison of sterile males and field-female killing systems. Theoretical and Applied Genetics 76: 870-879. Foster, G. G., G. L. Weller and G. M. Clarke, 1991 Male crossing over and genetic sexing systems in the Australian sheep blowfly Lucilia cuprina. Heredity 67: 365-371. Foster, J., M. Ganatra, I. Kamal, J. Ware, K. Makarova et al., 2005 The Wolbachia genome of Brugia malayi: endosymbiont evolution within a human pathogenic nematode. PLos biol 3: e121. Frisch, J., C. O’neill and M. Kelly, 2000 Using genetics to control cattle parasites—the Rockhampton experience. International Journal for Parasitology 30: 253-264. Frydman, H. M., J. M. Li, D. N. Robson and E. Wieschaus, 2006 Somatic stem cell niche tropism in Wolbachia. Nature 441: 509-512. Grenier, S., S. M. Gomes, B. Pintureau, F. Lasslbiere and P. Bolland, 2002 Use of tetracycline in larval diet to study the effect of Wolbachia on host fecundity and clarify taxonomic status of Trichogramma species in cured bisexual lines. J Invertebr Pathol 80: 13-21.
Page 39 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
Grenier, S., B. Pintureau, A. Heddi, F. Lassabliere, C. Jager et al., 1998 Successful horizontal transfer of Wolbachia symbionts between Trichogramma wasps. P Roy Soc B-Biol Sci 265: 1441-1445. Guglielmone, A. A., E. Gimeno, W. F. Idiart J Fisher, M. M. Volpogni, O. Quaino et al., 1999 Skin lesions and cattle hide damage from Haematobia irritans infestations. Medical and Veterinary Entomology 13: 324-329. Herbert, R. I., and E. A. McGraw, 2018 The nature of the immune response in novel Wolbachia-host associations. Symbiosis 74: 225-236. Hibler, C. P., 1966 Development of Stephanofilaria stilesi in the horn fly. J Parasitol 52: 890-898. Hill, J. K., H. M. Griffiths and C. D. Thomas, 2011 Climate change and evolutionary adaptations at species' range margins. Annual review of entomology 56: 143-159. Hoffmann, A. A., I. Iturbe-Ormaetxe, A. G. Callahan, B. Phillips, K. Billington et al., 2014 Stability of the wMel Wolbachia infection following invasion into Aedes aegypti populations. Plos Neglected tropical diseases 8: e3115 Hoffmann, A. A., B. Montgomery, J. Popovici, I. Iturbe-Ormaetxe, P. Johnson et al., 2011 Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature 476: 454. Hoffmann, A. A., P. A. Ross and G. Rašić, 2015 Wolbachia strains for disease control: ecological and evolutionary considerations. Evolutionary applications 8: 751-768. Hoffmann, A. A., and M. Turelli, 2013 Facilitating Wolbachia introductions into mosquito populations through insecticide-resistance selection. Proceedings of the Royal Society B: Biological Sciences 280: 20130371. Hornok, S., G. Foldvari, V. Elek, V. Naranjo, R. Farkas et al., 2008 Molecular identification of Anaplasma marginale and rickettsial endosymbionts in blood-sucking flies (Diptera: Tabanidae, Muscidae) and hard ticks (Acari: Ixodidae). Vet Parasitol 154: 354-359. Hughes, G. L., R. Koga, P. Xue, T. Fukatsu and J. L. Rasgon, 2011 Wolbachia infections are virulent and inhibit the human malaria parasite Plasmodium falciparum in Anopheles gambiae. PLoS Pathog 7: e1002043. Hughes, G. L., and J. L. Rasgon, 2014 Transinfection: a method to investigate Wolbachia–host interactions and control arthropod‐borne disease. Insect Mol Biol 23: 141-151. Iturbe‐Ormaetxe, I., T. Walker and S. LO'Neill, 2011 Wolbachia and the biological control of mosquito‐borne disease. EMBO reports 12: 508-518. Iwasa, M., and N. Ishiguro, 2010 Genetic and morphological differences of Haematobia irritans and H. exigua, and molecular phylogeny of Japanese Stomoxyini flies (Diptera, Muscidae). Med. Entomol. Zool. 61: 335-344. James, P., 2013 In vitro culture of buffalo fly and infections with Wolbachia. Jeyaprakash, A., and M. A. Hoy, 2000 Long PCR improves Wolbachia DNA amplification: wsp sequences found in 76% of sixty‐three arthropod species. Insect Mol Biol 9: 393-405. Johnson, S., R. Arthur and R. Shepherd, 1986 The distribution and prevalence of stephanofilariasis in cattle in Queensland. Australian Veterinary Journal 63: 121-124. Jones, S. R., and S. E. Kunz, 1996 Effects of immersion in water on survival of preimaginal stages of Haematobia irritans (Diptera: Muscidae). Journal of medical entomology 33: 27-31. Jonsson, N., and A. Matschoss, 1998 Attitudes and practices of Queensland dairy farmers to the control of the cattle tick, Boophilus microplus. Australian Veterinary Journal 76: 746-751. Jonsson, N. N., and D. G. Mayer, 1999 Estimation of the effects of buffalo fly (Haematobia irritans
Page 40 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
exigua) on the milk production of dairy cattle based on a meta-analysis of literature data. Med Vet Entomol 13: 372-376. Joshi, D., M. J. McFadden, D. Bevins, F. Zhang and Z. Xi, 2014 Wolbachia strain wAlbB confers both fitness costs and benefit on Anopheles stephensi. Parasit Vectors 7: 336. Kambris, Z., P. E. Cook, H. K. Phuc and S. P. Sinkins, 2009 Immune activation by life-shortening Wolbachia and reduced filarial competence in mosquitoes. Science 326: 134-136. Kang, L., X. Ma, L. Cai, S. Liao, L. Sun et al., 2003 Superinfection of Laodelphax striatellus with Wolbachia from Drosophila simulans. Heredity (Edinb) 90: 71-76. Kittayapong, P., S. Ninphanomchai, W. Limohpasmanee, C. Chansang, U. Chansang et al., 2019 Combined sterile insect technique and incompatible insect technique: The first proof-of-concept to suppress Aedes aegypti vector populations in semi-rural settings in Thailand. PLoS Negl Trop Dis 13: e0007771. Knipling, E., 1972 Entomology and the management of man's environment. Australian Journal of Entomology 11: 153-167. Knipling, E. F., 1972 Integrated control of livestock insect pests. Toxicology, Biodegradation and Efficacy of Livestock Pesticides. Koga, R., T. Tsuchida and T. Fukatsu, 2009 Quenching autofluorescence of insect tissues for in situ detection of endosymbionts. Appl Entomol Zool 44: 281-291. Kusmintarsih, E. S., 2009 Horizontal Transfer of the “Popcorn-Effect” Strain of Wolbachia from Drosophila melanogaster to Stomoxys calcitrans. Microbiology Indonesia 3: 121-125 Lane, J., T. Jubb, R. Shephard, J. Webb-Ware and G. Fordyce, 2015 Priority list of endemic diseases for the red meat industries. Meat & Livestock Australia Limited Project code B. AHE.0010. Laven, H., 1967 Eradication of Culex Pipiens Fatigans through Cytoplasmic Incompatibility. Nature 216: 383-384. Madhav, M., G. Brown, J. A. Morgan, S. Asgari, E. A. McGraw et al., 2020 Wolbachia successfully replicate in a newly established horn fly, Haematobia irritans irritans (L.) (Diptera: Muscidae) cell line. Pest Manag Sci 76:2441-2552 Madhav, M., R. Parry, J. A. Morgan, P. James and S. Asgari, 2020 Wolbachia endosymbiont of the horn fly (Haematobia irritans irritans): a Supergroup A strain with multiple horizontally acquired cytoplasmic incompatibility genes. Applied and Environmental Microbiology 86: e02589–02519 Mains, J. W., C. L. Brelsfoard, R. I. Rose and S. L. Dobson, 2016 Female adult Aedes albopictus suppression by Wolbachia-infected male mosquitoes. Scientific reports 6: 33846. Mains, J. W., P. H. Kelly, K. L. Dobson, W. D. Petrie and S. L. Dobson, 2019 Localized control of Aedes aegypti (Diptera: Culicidae) in Miami, FL, via inundative releases of Wolbachia-infected male mosquitoes. J Med Entomol 56: 1296-1303. Mateos, M., H. Martinez Montoya, S. B. Lanzavecchia, C. Conte, K. Guillén et al., 2020 Wolbachia pipientis associated with tephritid fruit fly pests: from basic research to applications. Frontiers in Microbiology 11: 1080. McGraw, E. A., and S. L. O'Neill, 2013 Beyond insecticides: new thinking on an ancient problem. Nature Reviews Microbiology 11: 181-193. McMeniman, C. J., 2009 Generation and characterization of a life-shortening Wolbachia infection in the dengue vector Aedes aegypti., PhD thesis. University of Queensland. 250 pp. McMeniman, C. J., A. M. Lane, A. W. Fong, D. A. Voronin, I. Iturbe-Ormaetxe et al., 2008 Host
Page 41 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
adaptation of a Wolbachia strain after long-term serial passage in mosquito cell lines. Applied and environmental microbiology. Applied and environmental microbiology 74: 6963-6969. McMeniman, C. J., R. V. Lane, B. N. Cass, A. W. Fong, M. Sidhu et al., 2009 Stable introduction of a life-shortening Wolbachia infection into the mosquito Aedes aegypti. Science 323: 141-144. McMeniman, C. J., and S. L. O'Neill, 2010 A virulent Wolbachia infection decreases the viability of the dengue vector Aedes aegypti during periods of embryonic quiescence. PLoS Negl Trop Dis 4: e748. Mendes, J., and A. Linhares, 1999 Diapause, pupation sites and parasitism of the horn fly, Haematobia irritans, in south‐eastern Brazil. Medical and Veterinary Entomology 13: 185-190. Min, K.-T., and S. Benzer, 1997 Wolbachia, normally a symbiont of Drosophila, can be virulent, causing degeneration and early death. Proceedings of the National Academy of Sciences 94: 10792- 10796. Moreira, L. A., I. Iturbe-Ormaetxe, J. A. Jeffery, G. Lu, A. T. Pyke et al., 2009 A Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya, and Plasmodium. Cell 139: 1268-1278. Moretti, R., and M. Calvitti, 2013 Male mating performance and cytoplasmic incompatibility in a wPip Wolbachia trans-infected line of Aedes albopictus (Stegomyia albopicta). Med Vet Entomol 27: 377-386. Nazni, W. A., A. A. Hoffmann, A. NoorAfizah, Y. L. Cheong, M. V. Mancini et al., 2019 Establishment of Wolbachia Strain wAlbB in Malaysian populations of Aedes aegypti for Dengue control. Curr Biol 29: 4241-4248 e4245. Negri, I., 2012 Wolbachia as an “infectious” extrinsic factor manipulating host signaling pathways. Front Endocrino 2: 115. O'Connor, L., C. Plichart, A. C. Sang, C. L. Brelsfoard, H. C. Bossin et al., 2012 Open release of male mosquitoes infected with a wolbachia biopesticide: field performance and infection containment. PLoS Negl Trop Dis 6: e1797. O'Neill, S. L., P. A. Ryan, A. P. Turley, G. Wilson, K. Retzki et al., 2018 Scaled deployment of Wolbachia to protect the community from dengue and other Aedes transmitted arboviruses. Gates Open Res 2: 36. Oyarzún, M., A. Quiroz and M. Birkett, 2008 Insecticide resistance in the horn fly: alternative control strategies. Medical and veterinary entomology 22: 188-202. Palavesam, A., F. D. Guerrero, A. M. Heekin, J. Wang, S. E. Dowd et al., 2012 Pyrosequencing-based analysis of the microbiome associated with the horn fly, Haematobia irritans. PLoS One 7: e44390. Parliament of Queensland. 1965. Buffalo Fly Control Acts, 1941 to 1965. Office of the Queensland Parliamentary Counsel, _https://media.sclqld.org.au/documents/digitisation/v16_pp507- 512_Stock_Buffalo%20Fly%20Control%20Acts,%201941%20to%201965.pdf Peters, T. M., and P. Barbosa, 1977 Influence of Population-Density on Size, Fecundity, and Developmental Rate of Insects in Culture. Annu Rev Entomol 22: 431-450. Pietri, J. E., H. DeBruhl and W. Sullivan, 2016 The rich somatic life of Wolbachia. Microbiology open 5: 923-936. Rigaud, T., J. Moreau and P. Juchault, 1999 Wolbachia infection in the terrestrial isopod oniscus asellus: sex ratio distortion and effect on fecundity. Heredity 83: 469-475. Ritchie, S. A., 2014 Rear and release: a new paradigm for dengue control. Austral Entomology 53: 363-367. Ritchie, S. A., M. Townsend, C. J. Paton, A. G. Callahan and A. A. Hoffmann, 2015 Application of
Page 42 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
wMelPop Wolbachia Strain to Crash Local Populations of Aedes aegypti. PLoS Negl Trop Dis 9: e0003930. Ross, P. A., N. M. Endersby and A. A. Hoffmann, 2015 Substantial fitness costs for Wolbachia infection on the starvation resistance of Aedes aegypti larvae, pp. Am J Trop Med Hyg. 93: 216-216 Ross, P. A., N. M. Endersby, H. L. Yeap and A. A. Hoffmann, 2014 Larval competition extends developmental time and decreases adult size of wMelPop Wolbachia-infected Aedes aegypti. Am J Trop Med Hyg 91: 198-205. Rothwell, J. T., J. A. Morgan, P. J. James, G. W. Brown, F. D. Guerrero et al., 2011 Mechanism of resistance to synthetic pyrethroids in buffalo flies in south-east Queensland. Aust Vet J 89: 70-72. Ruang-Areerate, T., and P. Kittayapong, 2006 Wolbachia transinfection in Aedes aegypti: a potential gene driver of dengue vectors. Proc Natl Acad Sci U S A 103: 12534-12539. Sackett, D., Holmes, P., Abbott, K., Jephcott, S., Barber, M., 2006 Assessing the economic cost of endemic disease on the profitability of Australian beef cattle and sheep producers. Final report of project AHW-087 Meat and Livestock Australia, Sydney, 119 pp Sarakatsanou, A., A. D. Diamantidis, S. A. Papanastasiou, K. Bourtzis and N. T. Papadopoulos, 2011 Effects of Wolbachia on fitness of the Mediterranean fruit fly (Diptera: Tephritidae). J Appl Entomol 135: 554-563. Schmidt, T. L., N. H. Barton, G. Rašić, A. P. Turley, B. L. Montgomery et al., 2017 Local introduction and heterogeneous spatial spread of dengue-suppressing Wolbachia through an urban population of Aedes aegypti. PLoS biology 15: e2001894. Schmittgen, T. D., and K. J. Livak, 2008 Analyzing real-time PCR data by the comparative C(T) method. Nat Protoc 3: 1101-1108. Schnitzerling, H. J., P. J. Noble, A. Macqueen and R. J. Dunham, 1982 Resistance of the buffalo fly, Haematobia irritans exigua (De Meijere), to two synthetic pyrethroids and DDT. .Australian Journal of Entomology 21: 77-80. Scott, M. J., 2014 Development and evaluation of male-only strains of the Australian sheep blowfly, Lucilia cuprina. BMC genetics 15: S3. Seawright, J., B. Birky and B. Smittle, 1986 Use of a genetic technique for separating the sexes of the stable fly (Diptera: Muscidae). Journal of economic entomology 79: 1413-1417. Seddon, H. R., 1967 Diseases of domestic animals in Australia. 2. Arthropod infestations. Serv. PubZs Dept HZth Aust. No. 6. Shaw, S. A., and I. A. Sutherland, 2006 The prevalence of Stephanofilaria sp in buffalo fly, Haematobia irritans exigua, in Central Queensland. Australian Journal of Entomology 45: 198-201. Showler, A. T., W. L. Osbrink and K. H. Lohmeyer, 2014 Horn fly, Haematobia irritans irritans (L.), overwintering. International Journal of Insect Science 6: IJIS. S15246. Snyder, F. M. 1965. Diptera. Muscidae. Insects of Micronesia 13: 191-327. Stegeman, D.A., Tozer, R.S., Sutherst R.W., 1996 Procedures for mass rearing the buffalo fly, Haematobia irritans exigua de Meijere (Diptera: Muscidae). Australian Journal of Entomology 35: 77- 79. Sutherst, R., A. Bourne, G. Maywald and G. Seifert, 2006 Prevalence, severity, and heritability of Stephanofilaria lesions on cattle in central and southern Queensland, Australia. Australian journal of agricultural research 57: 743-750. Sutherst, R. W., and R. S. Tozer, 1995 Control of Buffalo Fly (Haematobia-Irritans Exigua De Meijere)
Page 43 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
on Dairy and Beef-Cattle Using Traps. Aust J Agr Res 46: 269-284. Tillyard, R., 1931 The Buffalo-fly in Australia. Journal of the Council for Scientific and Industrial Research, Australia 4. Torres, L., C. Almazan, N. Ayllon, R. C. Galindo, R. Rosario-Cruz et al., 2012 Identification of microorganisms in partially fed female horn flies, Haematobia irritans. Parasitol Res 111: 1391-1395. Turley, A. P., L. A. Moreira, S. L. O'Neill and E. A. McGraw, 2009 Wolbachia infection reduces blood- feeding success in the dengue fever mosquito, Aedes aegypti. PLoS Neglected Tropical Diseases 3: e516. Urech, R., G. W. Brown, C. J. Moore and P. E. Green, 2005 Cuticular hydrocarbons of buffalo fly, Haematobia exigua, and chemotaxonomic differentiation from horn fly, H. irritans. J Chem Ecol 31: 2451-2461. Vreysen, M., A. Robinson, J. Hendrichs and P. Kenmore, 2007 Area-wide integrated pest management (AW-IPM): principles, practice and prospects, pp. 3-33 in Area-wide control of insect pests. Springer. Vreysen, M., K. Saleh, M. Ali, A. Abdulla, Z. Zhu et al., 2000 The use of the sterile insect technique (SIT) for the eradication of the tsetse fly Glossina austeni (Diptera: Glossinidae) on the Island of Unguja (Zanzibar). Journal of Economic Entomology 93: 123-135. Vreysen, M. J., and A. S. Robinson, 2011 Ionising radiation and area-wide management of insect pests to promote sustainable agriculture, pp. 671-692 in Sustainable Agriculture Volume 2. Springer. Vreysen, M. J., K. Saleh, F. Mramba, A. Parker, U. Feldmann et al., 2014 Sterile insects to enhance agricultural development: the case of sustainable tsetse eradication on Unguja Island, Zanzibar, using an area-wide integrated pest management approach. PLoS neglected tropical diseases 8: e2857. Vreysen, M. J., M. T. Seck, B. Sall and J. Bouyer, 2013 Tsetse flies: their biology and control using area-wide integrated pest management approaches. Journal of invertebrate pathology 112: S15-S25. Walker, T., P. Johnson, L. Moreira, I. Iturbe-Ormaetxe, F. Frentiu et al., 2011 The wMel Wolbachia strain blocks dengue and invades caged Aedes aegypti populations. Nature 476: 450-453. Weeks, A. R., M. Turelli, W. R. Harcombe, K. T. Reynolds and A. A. Hoffmann, 2007 From parasite to mutualist: rapid evolution of Wolbachia in natural populations of Drosophila. PLoS Biol 5: e114. Williams, J. D., R. W. Sutherst, G. F. Maywald and C. T. Petherbridge, 1985 The southward spread of buffalo fly (Haematobia irritans exigua) in eastern Australia and its survival through a severe winter. Aust Vet J 62: 367-369. Xu, Q., F. D. Guerrero, A. Palavesam and A. A. Perez de Leon, 2016 Use of electroporation as an option to transform the horn fly, Haematobia irritans: a species recalcitrant to microinjection. Insect Sci 23: 621-629. Yeap, H. L., J. K. Axford, J. Popovici, N. M. Endersby, I. Iturbe-Ormaetxe et al., 2014 Assessing quality of life-shortening Wolbachia-infected Aedes aegypti mosquitoes in the field based on capture rates and morphometric assessments. Parasites and vectors 7: 58. Yeap, H. L., P. Mee, T. Walker, A. R. Weeks, S. L. O'Neill et al., 2011 Dynamics of the “popcorn” Wolbachia infection in outbred Aedes aegypti informs prospects for mosquito vector control. Genetics 187: 583-595. Zabalou, S., A. Apostolaki, I. Livadaras, G. Franz, A. Robinson et al., 2009 Incompatible insect technique: incompatible males from a Ceratitis capitata genetic sexing strain. Entomologia Experimentalis et Applicata 132: 232-240.
Page 44 of 45
B.AHE.0242 – Wolbachia control of buffalo fly
Zhang, B., E. McGraw, K. D. Floate, P. James, W. Jorgensen et al., 2009 Wolbachia infection in Australasian and North American populations of Haematobia irritans (Diptera: Muscidae). Veterinary parasitology 162: 350-353. Zhang, D., R. S. Lees, Z. Xi, J. R. Gilles and K. Bourtzis, 2015 Combining the sterile insect technique with Wolbachia-based approaches: II-a safer approach to Aedes albopictus population suppression programmes, designed to minimize the consequences of inadvertent female release. PloS one 10: e0135194. Zheng, X., D. Zhang, Y. Li, C. Yang, Y. Wu et al., 2019 Incompatible and sterile insect techniques combined eliminate mosquitoes. Nature 572: 56-61.
Zug, R., and P. Hammerstein, 2012 Still a host of hosts for Wolbachia: analysis of recent data suggests that 40% of terrestrial arthropod species are infected. PloS one 7: e38544.
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