Chapter 1 Literature Review1

Chapter 1

Literature review1

1.1 Introduction

Australian mosquitoes of the genera Aedes (Kay 1982, Watson and Kay 1998, Boyd and Kay 1999, Watson and Kay 1999b), Culex (Boyd and Kay 2000) and Verrallina (Ryan et al. 2000) have been implicated in the transmission of over 500 arboviruses, including: Ross River, Barmah Forest and dengue viruses. Since 1991, over 77,000 human infections with these viruses have been reported in Australia (Communicable Diseases Network – Australia New Zealand – National Notifiable Diseases Surveillance System), leading to considerable public health concern. As there are no vaccines against these viruses, except for Japanese encephalitis, the community relies on personal protection against mosquito exposure, and on mosquito control to minimise the transmission of disease (Russell 1998a, Russell and Kay 2004).

The primary focus of mosquito control in Australia is reducing larval populations and is undertaken by local government authorities in Queensland, but also by State authorities, to varying degrees, in the remaining states. Successful larval control has involved the integration of a number of techniques including: • Public education: the community is urged to remove containers holding water from household gardens and to use personal protection; applicable to Aedes aegypti (Linnaeus) and Aedes notoscriptus (Skuse).

1 Aspects of this review, regarding the use of Bacillus thuringiensis var. israelensis for the control of immature mosquitoes, have been published in the proceedings of the 5th Pacific Rim conference on the biotechnology of Bacillus thuringiensis, pp 57 – 72 (Appendix A). 2

• Physical modification of habitats (runnelling): involves digging shallow channels to link isolated pools with the tidal source, this process increases the movement of low amplitude tides, regularly flushing the saltmarsh (Hulsman et al. 1989); widespread use against Aedes vigilax and some use against Aedes camptorhynchus (Thomson). • Biological control: involves introducing predators (e.g. copepods and fish) into larval habitats (Hurst 2004, Kay and Nam 2005); used primarily against Ae. aegypti, but native fish are applied to constructed wetlands against freshwater species, mainly Culex annulirostris. • Insecticides: include microbials and insect growth regulators, which have been used for the global control of mosquitoes for more than 20 years (Mian and Mulla 1982, Margalit and Dean 1985); broadly used for saltmarsh/mangrove and freshwater species.

This thesis will focus on improving the efficacy of insecticide use, while minimising environmental impacts. As such, the following review outlines: the significance of vector- borne disease in Australia, the habitats used by mosquito larvae, microbial insecticide and insect growth regulator use, and possible environmental impacts.

1.2 Mosquito-borne disease in Australia

Mosquitoes transmit three subgroups of pathogens: 1) filaria, 2) malaria protozoans, and 3) arboviruses. Human filariasis is no longer a medical concern in Australia; it was eradicated in the 1950s (Boreham and Marks 1986). Malaria was eradicated in 1981; however, up to 1,000 cases are imported annually by infected travellers, occasionally resulting in local transmission (Brookes et al. 1997, Russell 1998a). Arboviruses are of primary concern in Australia. Human disease results from infection with Ross River, Barmah Forest, dengue, Murray Valley encephalitis, Kunjin and Japanese encephalitis viruses (Table 1.1).

Ross River and Barmah Forest viruses are closely related Alphaviruses, and can lead to the onset of debilitating polyarthritis (Kay and Aaskov 1989, Mackenzie et al. 1998, Russell and Kay 2004). The severity and duration of symptoms due to Ross River and Barmah

Literature review 3 Victoria; TAS = ley encephalitis virus; JEV = , Dale et al. 1986, Ritchie et al. et al. 2000 and Kay 1999 1997b, Boyd and Kay 1999 Ritchie et al. 1997b, Boyd and Kay 2000, Ryan Reference Jeffery et al. 2002, Kay and Standfast 1987 Ritchie et al. 1997b, Ryan et al. 2000, Ryan Watson and Kay 1999b, Knox et al. 2003 Ritchie et al. 1997b, Watson and Kay 1998 Russell 1998a Campbell et al. 1989, Kay and Standfast 1987 Ritchie et al. 1997b Ritchie et al. 1997b, Ryan et al. 2000 b JEV, KUNV Associated arboviruses RRV, BFV RRV, BFV DENV RRV squitoes and the associated human diseases. a n Australia; NSW = New South Wales; SA = South Australia; VIC = ; DENV = Dengue virus; KUNV = Kunjin virus; MVEV = Murray Val Distribution All states/territories RRV, BFV, MVEV, QLD, NSW QLD, NSW, VIC Nth QLD All states/territories RRV All states/territories Nuisance QLD,NSW,NT,WA,SA RRV,BFV,MVEV Kay 1982 NSW, TAS, SA, VIC, WA RRV, BFV QLD,NSW,NT,WA RRV QLD, NSW, NT Cx. annulirostris Ae. procax Cq. linealis Ae. aegypti Ae. notoscriptus Cx. quinquefasciatus Ae. vigilax Ae. camptorhynchus Cx. sitiens Ve. funerea Both Ground-pool Container Japanese encephalitis virus. Tasmania. b. RRV = Ross River virus; BFV = Barmah Forest virus Notes a. NT = Northern Territory; QLD = Queensland; WA = Wester LarvalhabitatFreshwater Species Saltmarsh/mangrove Table 1.1: The larval habitats and distribution of Australian mo

Forest virus infection vary greatly among patients and are usually resolved in three months (Mackenzie and Smith 1996, Flexman et al. 1998, Harley et al. 2001).

On average 4,800 cases of Ross River virus are reported per year (Miller et al. 2003, Kelly- Hope et al. 2004). Ross River virus has been isolated from wild caught mosquitoes in most regions of Australia. The main vectors are: Ae. camptorhynchus (Campbell et al. 1989), Ae. vigilax (Kay 1982, Ryan et al. 2000), Ae. notoscriptus (Watson and Kay 1998), Cx. annulirostris (Ryan et al 2000), Verrallina funerea (Theobald) (Ryan et al. 2000), Aedes procax (Skuse) (Ritchie et al. 1997b, Ryan et al. 2000), and Coquillettidia linealis (Skuse) (Jeffery et al. 2002). The differing geographical distribution of each vector leads to areas of regional transmission.

There are over 700 human infections with Barmah Forest virus reported per year (Miller et al. 2003). The potential vectors of Barmah Forest virus are: Cx. annulirostris (Boyd and Kay 2000), Ae. camptorhynchus (Kay and Standfast 1987), Ae. vigilax (Boyd and Kay 1999), Ae. procax (Ryan and Kay 1999), and Cq. linealis (Kay and Standfast 1987).

Infection with dengue can lead to a non-specific febrile illness in most patients, ranging up to a fatal hemorrhagic disease (Gubler 1998b). Aedes aegypti is the major recognised dengue vector on mainland Australia (Watson and Kay 1999b, Knox et al. 2003), and is established in northern Queensland (Sinclair 1992). In addition, members of the Aedes scutellaris group (known dengue vectors), are established on the Torres Strait islands and in Cape York (Rosen et al. 1985, Russell 1998a). Over the past decade there have been almost annual outbreaks of dengue (10 to 3,000 cases per outbreak), each presumably started via infected travellers (Russell 1993, 1998a). In 2005, Aedes albopictus (Skuse) invaded the Torres Strait and threatens to colonise mainland Australia, further increasing the risk of dengue transmission (Russell et al. 2005).

Murray Valley encephalitis and Kunjin viruses are closely related flaviviruses, both causing severe fever and encephalitis, and with Murray Valley encephalitis some fatalities (Marshall 1988). The main vector of Murray Valley encephalitis and Kunjin virus is Cx. annulirostris (Kay et al. 1984), although other species such as Culex australicus Dobrotworsky & Drummond (Russell 1998a) and Aedes normanensis (Taylor) (Broom et al. 1989) may have subsidiary roles. Over the past decade, isolated cases of both Murray Valley encephalitis Literature review 5 and Kunjin have been recorded and confined to tropical Western Australia, the Northern Territory and north Queensland (Marshall 1988, Miller et al. 2003, Russell and Kay 2004). Recent research by Hall et al. (2003) has shown that Kunjin is a subtype of West Nile virus. West Nile virus was introduced into the USA in 1999 and to date has caused over 19,700 human infections and over 780 fatalities (Centres for Disease Control and Prevention – Division of Vector-Borne Infectious Diseases).

The first recognised outbreak of Japanese encephalitis in Australia occurred in 1995, when three clinical cases (two fatal) were reported at Badu Island in the Torres Strait (Hanna et al. 1996). A further two cases were reported in 1998, one in the Torres Strait and one on the mainland at the Mitchell River (Hanna et al. 1999). Virus isolations have been made from wild-caught Cx. annulirostris and Culex sitiens Wiedemann, indicating these species to be potential vectors in Australia (Ritchie et al. 1997a, Hanna et al. 1999, Johansen et al. 2001, van den Hurk et al. 2003, 2006). Laboratory trials have indicated that Culex gelidus Theobald, Culex quinquefasciatus Say and Ae. notoscriptus may also play roles in localised transmission (van den Hurk et al. 2003).

1.3 Habitats of mosquito larvae

In Australia, mosquito control has focused on reducing larval populations, rather than targeting adults which are more dispersed (WHO 1997). There are two broad classes of larval habitats: freshwater (ground-pool and container) and saltmarsh/mangroves. Each habitat type is colonised by the larvae of specific mosquito species, and mosquito control operations must be tailored to suit each habitat.

1.3.1 Freshwater

1.3.1.1 Ground-pool

In Australia, ground-pool freshwater habitats include permanent and temporary water bodies, e.g. lakes, swamps, streams, wheel ruts and dams. The main species found in these habitats are Cx. annulirostris and Cx. quinquefasciatus (Lee et al. 1989). Large numbers of Ae. procax, Aedes normanensis, Aedes sagax (Skuse), Aedes bancroftianus (Edwards), Aedes vittiger (Skuse) (Lee et al. 1984) and Cq. linealis (Lee et al. 1988) are also produced following rains.

The larvae of Cx. annulirostris, will inhabit almost any form of natural and artificial freshwater. This species inhabits permanent water during the dry season and rapidly exploits shallow vegetated pools, one day after rainfall (McDonald and Buchanan 1981, Dale and Morris 1996, Watson and Kay 1999a) as well as water associated with irrigation, rice-fields or sewage effluent disposal (Marks 1982, Kay et al. 1992). The larvae of Culex quinquefasciatus on the other hand, prefers to inhabit polluted or highly organic habitats (Marks 1982), as well as artificial containers (see below).

Effective broad-scale control of ground-pool freshwater larvae is difficult to achieve, due to the complex of species and larval habitat types involved, and this is confounded by a lack of research. Another confounding factor is the cost implications for local governments to produce a program comparable to that carried out in saltmarshes and mangroves. Currently Bti is the main product used for control and it is applied to temporary ponds and rainwater drainage areas (see section 1.5.1).

1.3.1.2 Container

The main species inhabiting containers (natural or artificial) holding freshwater water are Ae. aegypti (Lee et al. 1987), Ae. notoscriptus (Lee et al. 1984) and Cx. quinquefasciatus (Lee et al. 1989). Literature review 7

The larvae of Ae. aegypti (found only in north Queensland) have been recorded in artificial containers (rainwater tanks, roof gutters, pot plant bases, tyres and rubbish: Tun-Lin et al. 1995, Ritchie 1997, Montgomery and Ritchie 2002); natural containers (fallen palm fronds and bromeliad leaf axils: Ritchie and Broadsmith 1997); as well as subterranean sites (mine shafts, service pits and wells: Russell et al. 1996a, 1997, 2002). A main operational concern is identifying key containers that produce disproportionately high numbers of larvae (Tun- Lin et al. 1995), such as subterranean sites during the winter (Russell et al. 1997) and roof gutters during summer (Montgomery and Ritchie 2002).

Similarly, Ae. notoscriptus larvae have been recorded in a diverse range of container habitats, including artificial (Marks 1982, Watson 1998), natural (Fanning et al. 1997, Ritchie and Broadsmith 1997) and subterranean (Russell et al. 1997, Watson 1998). It has been noted that the species prefers organically rich water in semi-shaded or shaded positions. The key containers are discarded household items (garbage bins, buckets, kitchen items) year round, and garden accoutrements during winter (Watson 1998).

The control of larvae in container habitats has proved largely ineffective to date. This is due to the cryptic nature of larval habitats, the high level of labour required for control programs, and the lack of motivation of householders to reduce breeding sites. House to house inspections and health education are the main methods used to control dengue vectors. In north Queensland, where surveillance is carried out by Queensland Health, containers that cannot be emptied are treated with s-methoprene (Queensland Health 2000). Bti has a limited role in controlling container breeding species (see section 1.5.1). During dengue epidemics the adults of Ae. aegypti are targeted with residual indoor sprays (Queensland Health 2000).

1.3.2 Saltmarshes and mangroves

Australian estuaries consist of a mangrove flat dominated by vascular trees, with a landward saltmarsh dominated by herbs and low shrubs (Adam 1990, 1994). The mangrove flat is inundated on a regular, often daily, basis by high tides, whereas the saltmarsh is only inundated by the highest spring high tides (Adam 1994, Morrisey 1995). As the water drains from the wetlands with the receding tide, water remains in substrate depressions, forming

8 stagnant pools in the saltmarsh and upper mangrove. The pools are colonised by the larvae of a range of mosquito species, often at very high densities.

The primary species found in Queensland, New South Wales, Western Australia and the Northern Territory is Ae. vigilax (Dale et al. 1986, Kay and Jorgensen 1986) and throughout the remainder of the country is Ae. camptorhynchus (Lee et al. 1984). Other species that are commonly found in saltmarshes and mangroves are Cx. sitiens (Lee et al. 1989) and Ve. funerea (Lee et al. 1987).

Broad-scale mosquito control in saltmarshes and mangroves is undertaken in Queensland, Western Australia and the Northern Territory, and to a lesser extent in the remaining states. The protocols for control are well developed; for example, in south-east Queensland, rainfall greater than 100 mm over 24 h or a tide greater than 2.45 m is used to indicate larval hatching. During the summer, large areas of saltmarsh and the upper mangroves are treated via aerial or ground based operations, primarily with Bti. However, the treatment of heavily vegetated areas is still more problematic than open areas, because of poorer penetration of product.

1.4 Early mosquito control efforts

Prior to World War II, mosquito control in Australia was basic and larval control was conducted using petroleum oil and petrol (Cooling 1913, Bertram 1927). Since these times, mosquito control has evolved through three phases in Australia involving the use of: 1) organochlorine, 2) organophosphate and 3) biologically-based insecticides.

From World War II to the late 1960s, organochlorines (dichloro-diphenyl-trichloroethane [DDT] and dieldrin) were used. Organochlorines were applied using backpack sprayers, which was a labour intensive method. Blanket treatments of suspected larval habitats were made and little effort was put into preliminary surveys (Lyons and Lee 1985). No procedures had been developed to assess the efficacy of treatments, and general observations sometimes indicated poor results, particularly in thickly vegetated areas Literature review 9

(Mackerras et al. 1950, Allen 1979). Organochlorines were observed to affect aquatic insects other than mosquitoes, suggesting impacts on the ecosystem and this was highlighted by Rachael Carson’s publication Silent Spring (1962). Furthermore, resistance of mosquito populations to organochlorines had been observed overseas (Brown and Pal 1971), raising concerns for Australian operations.

By the late 1960s, it became apparent that organophosphate-based insecticides were more effective for mosquito control and “safer” to other forms of aquatic life (Carson 1962). In a decade, the use of DDT was banned in the USA and many European countries, after it was reported to be toxic to wildlife and to bioaccumulate in the food chain (Carson 1962). A total ban on the use of DDT in Australia was introduced in 1987 (Department of Environment and Heritage 1997).

The organophosphates currently registered for broad-scale mosquito control in Australia are temephos, malathion and chlorpyrifos and in Queensland pirimiphos-methyl. Temephos, the most popular organophosphate, was initially applied using hand-held equipment (Abate 1SG: Standfast et al. 1970). This 1% sand granule was reformatted into a 5% granule to improve aerial treatment efficiency, as this formulation, Abate 5 SG, which was subsequently relabelled as Abate 50 SG, could be applied at 1 lb/ac (1.12 kg/ha; Kay et al. 1973), and a total of 800 tonnes of Abate 50SG was applied aerially thereafter (Figure 1.1).

Organophosphates are reported to have a low mammalian toxicity, and acute human toxicity is rare (Derache 1977). However, organophosphates, e.g. chlorpyrifos, have been reported to bioaccumulate through the food chain (Varo et al. 2002). The organophosphates registered for mosquito control are broad spectrum insecticides and are highly toxic to a range of Arthropoda, including Insecta (Porter and Goimerac 1969, Hurlbert et al. 1972, Didia et al. 1975), Crustacea (Ludwig et al. 1968, Mortimer and Chapman 1995, Guzzella et al. 1997, Key et al. 1998) and Cladocera (Didia et al. 1975). The numerous studies that assessed the toxicity of organophosphates to non-target organisms are reviewed elsewhere (see Barron and Woodburn 1995, Australian and New Zealand Conservation Council 2000, National Registration Authority 2000).

Over time, techniques to monitor the efficacy of insecticide use were developed, which involved collecting larvae from targeted populations and using electrophoretic analysis to

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Seasons between 1984 - 2005

Figure 1.1: Insecticide use for mosquito control between 1984 and 2005 in south-east Queensland by CLAG members (Gold Coast City Council, Redlands Shire Council, Logan City Council and Tweed Shire Council; information collated from CLAG quarterly reports 1984 – 2005). Literature review 11 test esterase activity and bioassays to test insecticide susceptibility. Using these methods, tolerance of Ae. vigilax populations to temephos was identified in Redland Shire Council (Cousineau 1993). Organophosphates were phased out during the 1990s (Figure 1.1), when microbial insecticides and insect growth regulators became available.

1.5 Microbial insecticides

Bacillus thuringiensis var. israelensis (Bti), a soil dwelling bacterial species, has been used for mosquito control for more than 10 years in Australia (Figure 1.1) and for more than 20 years elsewhere (Mian and Mulla 1982, Margalit and Dean 1985). Another bacterial species, Bacillus sphaericus (Bs), was registered for mosquito control in Australia in 2005.

The activity of microbial insecticides is attributed to crystallised proteins of the bacterium, and with regard to Bs they are encapsulated in a spore (Baumann et al. 1987, Federici et al. 1990). When the proteins are ingested by mosquito larvae, they are dissolved and activated by the alkaline pH and proteinases of the larval midgut. This releases endotoxins (4 from Bti and 2 binary toxins from Bs) that bind to receptors on the epithelial midgut cells. This results in an osmotic imbalance across the epithelial cell membranes leading to swelling and lysis of the affected cells, neutralising the alkalinity of the midgut, and causing insect death (Federici et al. 1990, Gill et al. 1992, Darboux et al. 2001).

Many factors influence the efficacy of microbial insecticide treatments, especially the feeding rate of larvae (as the proteins must be ingested), which can be influenced by: 1) larval age, 2) water temperature and 3) density of larvae. Firstly, the larvae are most susceptible as younger instars, due to the low feeding activity of mature fourth instars. For this same reason, the non-feeding pupal stage is immune to Bti and Bs applications (van Essen and Hembree 1980, Mulla 1990, Yousten et al. 1992). Secondly, as water temperature decreases so does feeding activity. For example, Aedes vexans (Meigen) larvae are 10 times more sensitive to Bti at 25°C than at 5°C (Becker et al. 1992). Thirdly, as the density of larvae increases, competition for food increases, so Bti and Bs dosages should be increased (Becker et al. 1993).

Extrinsic environmental characteristics also influence treatment efficacy, particularly sunlight which degrades the protoxins (Burke et al. 1983, Becker et al. 1992, de Melo- Santos et al. 2001). For example, 3 times more Bti is required to cause 90% mortality of Ae. aegypti at 48 h in sunlight compared to shade (Becker et al. 1992). Also the residual activity of a Bti tablet was reduced by at least 20 days, in the presence of strong sunlight (de Melo- Santos et al. 2001). The presence of organic matter in the water column was found to reduce Bti activity by Mulla et al. (1990). During the trial, the same concentration of Bti resulted in 95% mortality of Cx. quinquefasciatus in clear water and no mortality in polluted water, where dense phytoplankton provided an alternative food source.

The majority of trials that assess either laboratory or field efficacy of insecticides follow similar methods (e.g. Brown et al. 1998b, Su and Mulla 1999, WHO 2005). However, it is difficult to directly compare results as a range of formulations and dosages have been used, and are often reported in different units: e.g. spores/ml; International Toxic Units [ITU]/ml or L; Aedes aegypti units/mg; ppm; or L/ha. Furthermore, if insecticide use is measured using surface area treated, the influence of water depth on concentration is not accounted for. Thus, the dimensions of water-bodies should be clearly stated or dosages presented as both a concentration of active ingredient (AI) (e.g. ppm AI) and a technical unit (e.g. L/ha).

1.5.1 Bacillus thuringiensis var. israelensis

Bti was initially described and identified as toxic to mosquito larvae by Goldberg and Margalit (1977). Since then, Bti-based products have been used for global mosquito control for more than 20 years. Bti has been used to control a range of mosquito genera including: Aedes (Goldberg and Margalit 1977, Becker and Rettich 1994), Anopheles (Goldberg and Margalit 1977, Amalraj et al. 2000, Fillinger et al. 2003), Culex (Goldberg and Margalit 1977, Becker and Rettich 1994, Su and Mulla 1999), Mansonia (Margalit and Dean 1985), and Coquillettidia (Margalit and Dean 1985) (Table 1.2).

Bti was initially registered for mosquito control in Australia in 1988. Early trials demonstrated that Australian mosquito larvae were susceptible to Bti (Davidson et al. 1981). However, Bti was slowly incorporated into control programs. This occurred as the necessary equipment to apply the products was not available, and aerial treatments of saltmarshes and Literature review 13 mangroves initially appeared to be 2 to 3 times more expensive than using temephos (Ritchie 1994). In spite of this, Bti is now the mainstay of mosquito control operations throughout Australia and extensive aerial applications are conducted.

It is widely accepted that Bti only provides short term control. The activity of the protoxins falls below acceptable levels for control after 2 to 3 days and is unable to be detected after 1 week (Karch et al. 1991, Kroeger et al. 1995, Fillinger et al. 2003). The residual activity of Bti is not increased by applying more product (Mulla et al. 1993).

Four formulations have been registered for use in Australia and the estimated field concentration of treatments can range from 250 to 1209 Bti ITU/L (Table 1.3). The most commonly used are the two liquid formulations (VectoBac 12AS and Teknar: 1,200 Bti ITU/mg), which are applied to saltmarsh/mangrove and ground-pool freshwater habitats using rotary-wing aircraft (1.2 L of Bti diluted in 2 L of water per ha; Woods and Dorr 1997, Muller et al. 2001). Small areas are treated using hand-held equipment, e.g. splatter guns often mounted on quad bikes. In south-east Queensland, upwards of 50,000 L are applied to saltmarshes and mangroves per year and 1,500 L are applied to ground-pool freshwater habitats per year (Contiguous Local Authorities Group [CLAG] 2005, North East Moreton Mosquito Organisation [NEMMO] 2005). Liquid Bti is reported to be effective against Ae. vigilax (e.g. Mottram et al. 1989, Brown et al. 2001, Muller et al. 2001) and Cx. sitiens (Brown et al. 1998b, Muller et al. 2001) in the laboratory and field, as well as Ae. camptorhynchus, Ae. notoscriptus (Brown et al. 2001), Ae. aegypti (Canyon 1997, Brown et al. 2001) and Ve. funerea (Jeffery et. al. 2005) in the laboratory (Table 1.2).

A new water dispersible formulation of Bti was registered for mosquito control in Australia in 2000. VectoBac WG (3,000 Bti ITU/mg) is transported and stored as micropellets, and once mixed with water it is applied as a liquid formulation (Abbott 1992). This formulation has advantages of economical transport, storage and long shelf life. However, application technology was only recently developed and VectoBac WG is not widely used (CLAG 2005, NEMMO 2005). Preliminary field trials found VectoBac WG to be as effective as the widely applied liquid VectoBac 12AS formulation for application against Ae. vigilax in saltmarshes and mangroves (Muller et al. 2001). Determination of product efficacy against the range of target species is required.

00 1999 Dame et al. 1981 Seleena et al. 2001 Amalraj et al. 2000 1 Riviere et al. 1987 1 2 1 Brown et al. 2001 1 11 Canyon 1997 Brown et al. 2001 1 Lee et al. 1996 1 Amalraj et al. 2000 1 Brown et al. 2001 1 1 Muller et al. 2001 2 Davidson et al. 1981 1 Mulla et al. 1990 1 Riviere et al. 1987 2 Skovmand and Sanogo 1 Davidson et al. 1981 2 Su and Mulla 1999 1 3 1 Lee et al. 1996 14 (Days) Reference Duration 0 % 50 50 95 50 50 80 100 100 Mortality c ITU/L) 71 72 50 100 55 120 388 774886 50 95 554 95 421 95 102 95 300 82 127 50 236 90 1800 90 1215 1019 95 1918 100 2100 90 1555 Dosage Bti ( b F 70000 100 14 Batra et al. 20 L F F L L L L F for the control of mosquito larvae. F.Polynesia L Australia L Malaysia L India Australia L USA Denmark L F.Polynesia L israelensis var. fluid Bacticide powder India Formulation Location Site Technical powder USA Bactimos Teknar VectoBac 12AS Australia L Teknar VectoBac 12AS Australia L VectoBac WG Australia L Comm. powder Australia F 1000 9 Bactimos Bti Comm. powder Australia L Commercial WG USA VectoBac 12AS India VectoBac 12AS Malaysia F 1918 Bacillus thuringiensis & a Ae. aegypti Ae. notoscriptus Cx. annulirostris Cx. annulirostris Cx. quinquefasciatus Cx. quinquefasciatus Species Freshwater larvae Table 1.2: Efficacy of Literature review 15 Ali et al. 2000 Muller et al. 2001 Brown et al. 1998b Jeffery et al. 2005 2 Karch et al. 1991 3 3 2 Su and Mulla 1999 2 Ritchie 1994 1 Ali et al. 1999 2 Mottram et al. 1989 11 Brownetal.2001 2 Webb and Russell 2001 2 Woods et al. 2001b 1 Ritchie 1994 1 2 2 1 2 2 2 1 Brownetal.2001 1 (Days) Reference Duration % 76 95 95 95 95 98 100 100 72-74 Mortality ratory data. If results could not be directly c ITU/L) 24 40 246 492 199 90 111421111 95 100 95 399 89.5 933 98.6 778 720 648 310 95 310 95 1269 100 1400 Dosage (Bti b F 5300 98 F 9620 99 F F 1555 95 F F F L F at are found in Australia. Bangladesh F USA Australia L Australia F Australia Australia Australia Australia Australia L ITU/L for ease of comparison between field and labo Bti VectoBac 12AS Zaire FormulationVectoBac G Location Site VectoBac TP Bangladesh L Bactimos pellets Australia F 1015 Teknar VectoBac 12AS Australia L VectoBac G VectoBac G VectoBac WG VectoBac 12ASVectoBac 12AS Australia L VectoBac 12AS Teknar VectoBac 12AS Australia L converted to ITU/L the paper was omitted from the table. Cx. sitiens Bti & a Ae. vigilax Ae. vigilax Ve. funerea Ae. camptorhynchus Cx. quinquefasciatus Cx. sitiens Values have been compared in Species Saltmarsh/mangrove larvae b. L = laboratory; F = field. c. Notes a. Examples of data are presented for only those species th Table 1.2 cont.

L L / / ITU ITU/L ITU Bti Bti Bti ITU/L b Bti 250 – 1000 518 – 1209 933 EFC a Registered treatment rate Label rate registered for mosquito control in Australia. ITU/mg 125–500g/ha ITU/mg 0.6–1.2L/ha 518–1036 ITU/mg 0.6–1.4L/ha a standard water depth of 15 cm for saltmarshes and ITU/mg 7kg/ha israelensis cers. Bti Bti Bti Bti . var. Activity Bacillus thuringiensis Aqueoussuspension 1200 Granular(corncob) 200 . c mangroves VectoBac G VectoBac WG Water dispersible granule 3000 Product NameTeknar Formulation VectoBac 12AS Aqueous suspension 1200 b. EFC = Estimated field concentration; calculated assuming c. Only registered for use in saltmarshes and mangroves Notes a. Recommended dosages according to the labels or produ Table 1.3: The formulations of Literature review 17

The granular formulation (VectoBac G: 200 Bti ITU/mg) is used in saltmarshes and mangroves where there is heavy vegetation and is not registered for use in freshwater habitats. It is applied via rotary-wing aircraft using under slung buckets or hoppers attached to the sides, or on the ground using quad bikes. VectoBac G has been recorded to control Ae. vigilax (Ritchie 1994) and Ve. funerea (Jeffery et al. 2005) in saltmarshes and mangroves. The low potency of VectoBac G means that large amounts of product must be carried to treat a small area. This leads to expensive treatments because extra ferrying time is required. However, there are no economical alternatives available. Granular s-methoprene (Altosid pellets), is sometimes used and although it is relatively expensive, it does provide residual control.

Bti is rarely used for the control of larvae in artificial containers (Ritchie 1997). Due to the cryptic nature of container habitats, operators rely on community participation for larval control. At present, the only product available for domestic use is granular s-methoprene, available at hardware stores. S-methoprene does not cause rapid death of larvae; rather it prevents pupae from emerging (see section 1.6.1). This means that to monitor product use the emergence of pupae must be recorded and this is something which the public would not do. Thus, Bti is seen as a more desirable product for community use and it has been effective against Ae. aegypti and Ae. notoscriptus in laboratory trials (Canyon 1997, Brown et al. 2001). No results from Australian field trials have been published. A handful of promising studies have been conducted overseas, where residual activity of Bti was recorded after 1 week in the field (Becker et al. 1991, Batra et al. 2000, Seleena et al. 2001, Mulla et al. 2004).

To date, no cases of larvae developing resistance to Bti have been recorded in the field (Becker and Ludwig 1993). The lack of resistance to Bti is most likely due to the complex of insecticidal proteins that are present. Selection trials in the laboratory have shown that the evolution of resistance to Bti is slow and correlated with the number of toxins present (Cheong et al. 1997). When the entire complex is present, only a low level of resistance develops (Georghiou and Wirth 1997).

Different operational procedures can be practiced in an effort to minimise the development of resistance to pesticides. The most popular method is to alternate between insecticides with different modes of action, in blocks of consecutive applications (Dame 2001). In south-

18 east Queensland, for example, Brown et al. (1998b) recommended that Bti should be rotated with the insect growth regulator, s-methoprene; with the latter being used early in the season in saltmarshes and mangroves, as it has a low efficacy against Cx. sitiens (Ritchie et al. 1997c), which become abundant from March onwards.

1.5.2 Bacillus sphaericus

Bs was initially described as moderately toxic to mosquito larvae by Kellen and Meyers (1964), and subsequently various researchers have isolated more than 150 strains that range from highly toxic to non-toxic. Bs exhibits the highest toxicity against larvae of the genus Culex (e.g. Karch et al. 1990, Brown et al. 2004) (Table 1.4), closely followed by the genus Anopheles (e.g. Lacey et al. 1988, Becker and Bozhao 1989, Barbazan et al. 1998).

The VectoLex formulation of Bs was registered for mosquito control in Australia in 2005. It is recommended for use in freshwater ground-pool habitats against Cx. annulirostris and Cx. quinquefasciatus. Both these species were recorded to be susceptible to Bs (VectoLex WDG: 650 Bs ITU/mg) in the laboratory and field by Brown et al. (2004). The Australian species that were not susceptible to Bs in the laboratory were Ae. aegypti, Ae. notoscriptus, Ae. vigilax (Brown et al. 2004) and Ve. funerea (Jeffery et al. 2005).

In contrast to Bti, the protoxins of Bs have been reported to have residual activity in the field. The residual activity of Bs (>90% mortality) was recorded to be up to 3 weeks in Australia (Brown et al. 2004); 4 weeks in the USA (Mulla et al. 1988, Matanmi et al. 1990); and 6 weeks in Africa (Nicolas et al. 1987). The efficacy and residual activity of Bs is directly correlated with the amount of product applied (Mulla et al. 1988). The residual activity of Bs is attributed to the spores replicating in larval cadavers and then being released into the water. Laboratory studies have indicated that the larval cadavers contain the necessary nutrients for the sporulation process and toxin synthesis (Davidson 1984, Des Rochers and Garcia 1984, Becker et al. 1995, Rives and Breitfuss 2005).

The resistance of larvae to Bs has been recorded during field studies in France (Chevillon et al. 2001), China (Yuan et al. 2000), India (Rao et al. 1995), Brazil (Silva-Filha et al. 1995), and Thailand (Mulla et al. 2003). The development of resistance to Bs is of greater concern Literature review 19 than Bti, as the potency is primarily attributed to a binary toxin binding to one specific receptor (Wirth et al 2000, Nielsen-LeRoux and Charles 1992). No cross resistance between Bti and Bs has been recorded (Yuan et al. 2003) and is unlikely to develop as the endotoxins, released after the proteins are broken down in the larval midgut, bind to different receptors on the epithelial midgut cells (Nielsen-LeRoux and Charles 1992, Rodcharoen and Mulla 1996).

1.6 Insect growth regulators

Insect growth regulators possess growth retarding properties and include juvenile hormone analogues. The first insect growth regulators developed were chemically related to the natural juvenile hormones of insects that regulate the moulting and metamorphosis of immature insects. The discovery that hormones controlled the growth and moulting of insects was first made in 1934 by Sir Vincent Wigglesworth (1934). The potential link between juvenile hormones and insect control was made by Williams (1956). This discovery lead to intensive research into the potential of growth hormones for insect control, resulting in over 500 juvenile hormone analogues being developed (Mian and Mulla 1982). Currently, s-methoprene is the only juvenile hormone analogue registered for mosquito control in Australia. An alternative juvenile hormone analogue, pyriproxyfen, is undergoing registration for mosquito control in Australia.

Unlike the natural juvenile hormones, the synthetic analogues are not affected by the destructive enzymes of the larval metabolism. The presence of the hormone mimic interferes with neuroendocrine functions and induces morphogenetic abnormalities and most commonly, pupae fail to emerge (Arias and Mulla 1975b, Gordon and Burford 1984, Sawby et al. 1992). Impacts on adults that emerge after exposure to sublethal doses of insect growth regulators have been observed, including: reduced longevity (Arias and Mulla 1975a, Robert and Olson 1989, Sawby et al. 1992), reduced biting activity (Ritchie et al. 1997c), unrotated genitalia, reduced fecundity (Arias and Mulla 1975a, Robert and Olson 1989) and reduced viability of eggs (Arias and Mulla 1975a).

Ali et al. 2000 Su and Mulla 1999 Floore et al. 2002 2 2 17 17 35 35 35 (Days) Reference Duration % 95 2 Brownetal.2004 95 2 90 90 50 2 Skovmand and Sanogo 1999 95 2 Brownetal.2004 67-97 78-98 67-100 50 – 100 81 – 100 Mortality c 3 2 2 ITU/L) 20 33 66 41 32 95 – 100 21 15 32 98 – 100 21 62 800 95 2 300 95 2 133 90 4 Ali et al. 1999 170 122 1048 95 2 Brown et al. 2004 1740 95 2 3077 66 – 100 14 Karch et al. 1991 <0.01 95 2 Dosage Bs ( b F F L L F F F Bangladesh F USA USA Denmark L Australia L USA F for the control of mosquito larvae. Liquid VectoLex CG Australia L VectoLex CG Australia L Formulation Location Site VectoLex WDG Australia L VectoLex CG Australia L VectoLex G Zaire F VectoLex WDG Australia L VectoLex CG Australia L VectoLex TP Bangladesh L VectoLex WDG Australia L VectoLex WDG Bacillus sphaericus a Ae. aegypti Ae. notoscriptus Cx. annulirostris Cx. quinquefasciatus Bs Species Freshwater larvae Table 1.4: Efficacy of Literature review 21 (Days) Reference Duration ata. If results could not be directly compared in % 95 2 95 2 Jeffery et al. 2005 95 2 Mortality c 5 ITU/L) 10 1835 95 2 Brown et al. 2004 1000 95 2 Dosage Bs ( b at are found in Australia. ITU/L for ease of comparison between field and laboratory d Bs VectoLex CG Australia L Formulation Location Site VectoLex WDG Australia L VectoLex CG Australia L VectoLex WDG Australia L VectoLex WDG Australia L >556000 a ITU/L the paper was omitted from the table. Ae. vigilax Ve. funerea Cx. sitiens Bs b. L = laboratory; F = field. c. Values have been converted to Notes a. Examples of data are presented for only those species th Species Saltmarsh/mangrove larvae Table 1.4 cont.

Considering that the activity of insect growth regulators works via ingestion or contact with the cuticle, they can be applied to either larvae or pupae. It has been noted that older, or fourth instar, larvae are the most susceptible (Schaefer and Wilder 1972, Estrada and Mulla 1986, Schaefer et al. 1988). Additionally, the activity of s-methoprene and pyriproxyfen is decreased in the presence of sunlight (Quistad et al. 1975, Schaefer et al. 1988) and organic matter in the water column (Schaefer and Dupras 1973, Schaefer et al. 1991).

1.6.1 s-methoprene

S-methoprene (isopropyl 11-methoxy-3,7,ll-trimethyl-2,4-dodecadienoate) has been used for mosquito control for more than 10 years in Australia and for more than 30 years overseas. Throughout the world, s-methoprene has been shown to be active against a range of mosquito genera, including: Aedes (Floore et al. 1991, Kramer et al. 1993, Ritchie et al. 1997c), Culex (Mulla and Darwazeh 1975, Floore et al. 1991), and Coquillettidia (Sjogren et al. 1986) (Table 1.5).

The resistance of natural populations to s-methoprene has been recorded in the field on two occasions. The first was as increased tolerance of an isolated population of Aedes taeniorhynchus (Wiedemann) on barrier islands in Florida that had been treated for 6 years with s-methoprene (Dame et al. 1998). The second was an increased tolerance of Aedes nigromaculis (Ludlow) in California that had been treated for 20 years with s-methoprene (Cornel et al. 2000). Many laboratory studies have induced s-methoprene resistance (Brown and Brown 1974, Brown et al. 1978). The low level of resistance developed in field populations is quite remarkable, considering that resistance to DDT and organophosphate based insecticides was detected within 3 years of introduction (Brown and Pal 1971).

Four formulations of s-methoprene are registered for use in Australia. The most widely applied is the sand formulation (Altosand: 4 g AI/kg). Altosand is useful for treating densely vegetated areas in ground-pool freshwater and saltmarsh/mangrove habitats. In south-east Queensland, between 1994 and 1998, the use of Altosand became popular when organophosphates (temephos) were phased out (Figure 1.1). During early trials by Ritchie (1994), Altosand was recorded to provide effective control of Ae. vigilax and medium control of Cx. sitiens. However, during 1997/98 results from a series of quality assurance Literature review 23 3 7 Jefferyetal.2005 7 Ritchie et al. 1997c 8 Ritchie 1994 7 Ritchie et al. 1997c 6 Brown et al. 2000a 8 Ritchie 1994 7 WHO 2000 77 Brown et al. 1999 Ritchie et al. 1997c 7 Brown et al. 2000b 7 7 Robert and Olson 1989 7 Ritchie et al. 1997c 7 Sawbyetal.1992 7 WHO 2000 7 Ritchie et al. 1997c 35 10 Ali et al. 1999 35 35 (Days) Reference Duration ould not be directly compared in ppb AI 98 95 90 90 90 71 90 95 90 98 50 95 90 90 90 50 97 90 100 inhibition % Emergence c 4 27 25 90 – 100 14 – 35 Self et al. 1978 52 50 10 1.2 31 – 99 140 Ritchie 1994 0.2 2.6 0.1 200 100 2.65 0.68 6.54 0.17 1.35 0.34 1.12 1.31 0.13 (ppb AI) Dosage b L L L F L Australia F Australia F 0.12 52 – 80 Australia L Australia L USA Australia F 0.12 66 – 100 Indonesia F Australia L USA Australia L Australia L USA Australia L comparison between field and laboratory data. If results c cies that are found in Australia. d Altosand Altosand ALL ALL Altosid pellets Australia F Liquid Technical grade Bangladesh L Altosand Altosid 10F ALL ALL ALL Liquid ALL Formulation Location Site Technical grade USA -methoprene for the control of mosquito larvae. s Cx. sitiens & a Ve. funerea Ae. vigilax Ae. vigilax Ae. notoscriptus Ae. aegypti Cx. sitiens Cx. quinquefasciatus Cx. annulirostris the paper was omitted from the table. Saltmarsh/mangrove larvae Species Freshwater larvae b. L = laboratory; F = field. c. Values have been converted to ppb AI for ease of Notes a. Examples of data are presented for only those spe d. ALL = Altosid Liquid Larvicide. Table 1.5: Efficacy of

24 trials indicated poor performance of Altosand (Brown and Kay 1999). This lead to an increased use of Bti. Altosand is slowly regaining popularity after formulation doubts were alleviated during a field trial by Brown et al. (2000b) and over 17,000 kg was used in the 2002/03 season by CLAG members, but only 6,120 kg was used in the 2004/05 season (Figure 1.1).

Liquid formulations (Altosid Liquid Larvicide: 50 g AI/L) of s-methoprene are also used for the control of ground-pool freshwater and saltmarsh/mangrove larvae. As liquid s- methoprene is relatively expensive compared with liquid Bti, it is generally viewed as a secondary tool: for use when adverse weather prevents treatments with Bti against younger instars (Mulla 1990), or as a resistance management tool (Dame 2001). The cost ratio of the minimum label rates for liquid Bti and s-methoprene is 1:4. Laboratory trials have demonstrated that the saltmarsh/mangrove species, Ae. vigilax (Ritchie et al. 1997c) and Ve. funerea (Ritchie et al. 1997c, Jeffery et al. 2005), are susceptible to liquid s-methoprene. Culex sitiens was less susceptible (Ritchie et al. 1997c) and treatment failures are likely to happen later in the season when this species is common in south-east Queensland. The freshwater species, Cx. annulirostris (Ritchie et al. 1997c, Brown et al. 2000b), Ae. aegypti and Ae. notoscriptus (Ritchie et al. 1997c) were also susceptible to s-methoprene in the laboratory.

The sand and liquid formulations of s-methoprene are active in the field for 7 to 10 days (Ritchie et al. 1997c, Brown et al. 2000b). The estimated field concentration of treatments ranges from 8 to 13 ppb AI (Table 1.6).

The sustained release s-methoprene formulations (Altosid Pellets: 40 g AI/kg and XR Briquets: 18 g AI/kg) remain active in the field for several months (Kramer and Beesley 1991, Kramer et al. 1993). They contain a high concentration of microencapsulated s- methoprene that is released over time and does not reach a concentration higher than 6 ppb AI in the water column (Ross et al. 1994a).

The sustained release briquets are designed for the control of mosquitoes in large bodies of water. They are impractical for use on saltmarsh/mangrove where water-bodies are scattered over a large area. Briquets are useful for control in artificial container habitats and ground- pool freshwater habitats. There are no peer-reviewed publications on the use of briquets in Literature review 25 c c b 8 – 13 ppb AI 219 – 438 ppb AI EFC or 88 ppb AI 80 – 320 ppb AI 2 a Registered treatment rate or 1 briquet/5000L 220–360ml/ha 7–12ppbAI a standard water depth of 15 cm for saltmarshes and cers. . 18 g AI/kg 1 briquet/10 – 20 m Activity Label rate 40 g AI/kg 3 – 4 kg/ha -methoprene registered for mosquito control in Australia. s Formulation Granular(sand) 4gAI/kg 3–5kg/ha Pellet mangroves. b. EFC = Estimated field concentration; calculated assuming c. Achieved only if all AI is released into water at one time Notes a. Recommended dosages according to the labels or produ Table 1.6: The formulations of Product Name Altosand Altosid liquid larvicideAltosid Liquid XR Concentrate briquets 50 g AI/L Briquet Altosid Pellets

Australia. This is not surprising as this product is cost prohibitive, retailing at around AUD $15 per briquet.

The sustained release pellet formulation (Altosid XRG: 40 g AI/kg) was designed for use in saltmarshes and mangroves and can be applied pre-flooding without any loss of activity. In trials in the USA this formulation was found to be effective in these habitats (Floore et al. 1990, 1991, Kramer and Beesley 1991, Kramer et al. 1993) and was first used commercially in Australia in 1997. The use of pellets in Australian saltmarshes and mangroves was studied by Ritchie (1994), who recorded effective control (>90% emergence inhibition) of Ae. vigilax for 3 months. This formulation has also been recorded to provide effective control (>90% emergence inhibition) of Ae. aegypti in containers for 6 months, in Australia (Ritchie and Broadsmith 1997). The use of pellets for control of ground-pool freshwater larvae in Australia has not been examined in any detail.

In New Zealand, a granular formulation of s-methoprene (ProLink XRG) is used in eradication programs on both the North and South islands against Ae. camptorhynchus which was imported from Australia in 1998. The strategy is based on maintaining a presence of s-methoprene in saltmarshes by reapplying every 21 d (Garner et al. 2001) and appears to have been effective with Ae. camptorhynchus eradicated from Hawkes Bay, Porangahau, Makia, Tairawhiti and Whitford, with ongoing programs at Kaipara, Wairau and Grassmere (South Island) (Frampton 2005) and, most recently in 2006, the Coromandel peninsula (B. Kay pers. comm.).

1.6.2 Pyriproxyfen

Pyriproxyfen (4-phenoxyphenyl (RS)-2-(2-pyridyloxy)propyl ether) is undergoing registration for mosquito control in Australia. No Australian studies on the efficacy of pyriproxyfen have been published. It has been used for mosquito control overseas for more than 15 years. Liquid and granular formulations have been developed and remain active in the field for up to 3 months (WHO 2000).

Throughout the world, pyriproxyfen has been effective for the control of Aedes, Anopheles and Culex spp. (Table 1.7). For example, in the USA pyriproxyfen was effective against Ae. Literature review 27 the Estrada and Mulla 1986 Ali et al. 1999 Mulla et al. 1986 Schaefer et al. 1988 42 60 Schaefer et al. 1991 42 14 14 10 10 14 Hatakoshi et al. 1987 14 Itoh 1994 77 Chavasse et al. 1995 14 14 14 14 14 Schaefer et al. 1988 68 Mulligan and Schaefer 1990 (Days) Reference Duration 50 95 50 90 50 50 50 90 50 95 100 100 100 inhibition % Emergence tory data. If results could not be directly compared in ppb AI c 50 50 2.6 1.1 0.3 0.16 0.006 100 0.018 0.005 100 (ppb AI) Dosage b L 20 99-100 42 WHO 2000 F 20 94-100 42 L 0.33 L 0.04 F 0.045 100 larvae. e found in Australia. USA USA Japan L 0.023 Tanzania F 100 USA comparison between field and labora Formulation Location Site GR S-31183 Technical grade Bangladesh L 0.29 Technical grade Japan L 0.023 EC S-31183 S-31183 EC USA a Ae. aegypti Cx. quinquefasciatus paper was omitted from the table. Species Freshwater larvae b. L = laboratory; F = field. c. Values have been converted to ppb AI for ease of Notes a. Examples of data are presented for only those species that ar Table 1.7: Efficacy of pyriproxyfen for the control of mosquito

28 aegypti in the laboratory (Estrada and Mulla 1986) and Cx. quinquefasciatus in the laboratory and the field for 2 months (Schaefer et al. 1988, Mulligan and Schaefer 1990). Pyriproxyfen was effective against Anopheles spp. for 2 months in the Solomon Islands (Okazawa et al. 1991) and in Sri Lanka (Yapabandara et al. 2001).

No reports of resistance to pyriproxyfen have been published. After 17 generations of pressure in the laboratory, Schaefer and Mulligan (1991) recorded no increased tolerance of Cx. quinquefasciatus to pyriproxyfen. The potential for cross-resistance for mosquito strains exposed to both s-methoprene and pyriproxyfen has not been explored.

1.7 Ecological impacts of biologically-based larvicides

A range of laboratory and field studies have been conducted to assess the impact of Bti and s-methoprene on non-target organisms. This review focuses on impacts in saltmarshes and mangroves, the primary target habitat of Australian mosquito control.

1.7.1 Bacillus thuringiensis var. israelensis

Much literature has been published about the impact of Bti on non-target organisms (Glare and O’Callaghan 1998b, Boisvert and Boisvert 2000, Stark 2005b) and the possible impacts on macroscopic organisms have been summarised in Table 1.8. The studies were conducted in different environments: 41% (n = 16) in the laboratory, 18% (n = 7) in mesocosms, 18% (n = 7) in saltmarshes/mangroves, and 23% (n = 9) in flowing rivers (black fly control). The impacts of normal operations were assessed by 69% of studies, and the rest (31%) were assessed as an overdose of Bti.

Bti does not appear to impact on Mollusca. No affects were recorded in field trials in saltmarshes in Australia (Barnes and Chapman 1998) or flowing rivers in the USA (Jackson et al. 1994, 2002). Additionally in the USA, Gastropods were not impacted in mesocosms (Physa sp.: Garcia et al. 1980) or lentic freshwater in the USA (Hershey et al. 1995, 1998), as were neither bivalves in lentic freshwater (Hershey et al. 1995, 1998). Literature review 29

Under laboratory conditions Bti is toxic to Chironomidae (Rey et al. 1998, Dickman 2000), but in the field, the possible impact is not clear. In flowing rivers, where black flies are the target, a moderate increase in drift of Chironomidae was been observed in some cases (Car and de Moor 1984, Back et al. 1985), but no effect was observed at other times (Colbo and Undeen 1980, Pistrang and Burger 1984, Merritt et al. 1989, Molloy 1992, Dickman 2000, Jackson et al. 2002). In streams, Bti is carried by the current and long-term exposure is unlikely to occur, which may be possible in the stagnant pools of mangroves and saltmarshes.

In mesocosms, a low level of toxicity of Bti to Chironomidae has been recorded (Ali 1981); yet, in another case no mortality was evident (Liber et al. 1998). In lentic freshwater, no impact was recorded by Hershey et al. (1995) in a small-scale field trial conducted over 4 months during which Bti was reapplied to three plots, nine times. Four benthic samples were analysed for each treated and untreated control plot after each addition of Bti. In contrast, during a long-term field trial by Hershey et al. (1998) and Niemi et al. (1999) a reduction in density was recorded after one year. During this study, nine lentic freshwater wetlands were treated with Bti (nine with s-methoprene) and nine remained untreated. The sites were treated at the highest recommended label rate and repeated every three weeks. During the first year of the study, a drought reduced insect numbers on both treatment and control sites. During the second and third years of the study, the lower number of Chironomidae on treated sites was attributed to their failure to recolonise. The results of Hershey et al. (1998) and Niemi et al. (1999) highlight the importance of field-based ecological trials.

Bti does not seem to affect other insect Families (aside from Chironomidae) at operational concentrations. In Australian saltmarshes, no impacts on insects were recorded by Barnes and Chapman (1998) during a well designed field trial. This study also used a before/after design, with adequate temporal sampling and results were compared with two reference locations, providing spatial replication of untreated controls.

The majority of trials have reported Bti to be non-toxic to Ephemeroptera. In trials conducted overseas, Ephemeroptera were unaffected by Bti exposure in the laboratory (Dickman 2000) and in mesocosms (Miura et al. 1980, Mulla et al. 1982, Ali 1981). In flowing rivers no effect was recorded by all (Colbo and Undeen 1980, Back et al. 1985, Car and de Moor 1984, Merritt et al. 1989, Molloy 1992, Jackson et al. 1994, 2002) but one case

2002 d Burger 1984 2003 Reference de Araujo-Coutinho et al. Barnes and Chapman 1998 Molloy 1992 c d Jackson et al. 2002 Duration b EFCEFC 3 y 3 y Hershey et al. 1998 Niemi et al. 1999 EFC 4 m Hershey et al. 1995 EFCEFC 4 m 3 y Hershey et al. 1995 Hershey et al. 1998 EFC 3 y Hershey et al. 1998 EFC 1 m Barnes and Chapman 1998 EFC N/S Garcia et al. 1980 EFC N/S Garcia et al. 1980 Dosage ) EFC 7 d Colbo and Undeen 1980 a F (B) EFC 3 d Car and de Moor 1984 F (B) EFC 6 d Merritt et al. 1989 Canada F (B) EFC 3 d Back et al. 1985 USA F (B) EFC 2 d Jackson et al. 1994, USA F (B) EFC 2 d Pistrang an USA L OD 4 d Reish et al. 1985 on non-target organisms. israelensis VectoBac G USA F (M) VectoBac 12AS USA F (B) EFC 2 Teknar Comm. formulation Brazil F (B) EFC 2 y VectoBac G USA F (M) Teknar VectoBac 12AS Australia F EFC 1 m VectoBac WP USA F (B) EFC 4 d VectoBac G USA F (M) var. Effect Formulation Location Site No impact VectoBac 12AS Australia F (M) No impact Comm. formulation Canada F (B No impact VectoBac G USA F (M) No impact Exp. formulation USA M No impact VectoBac G USA F (M) No impact Exp. formulation USA M 50% mortality Bactimos Reduced density Comm. formulation South Africa Reduced density VectoBac G USA F (M) Bacillus thuringiensis d sp. Chironomidae Mollusca Physa Bivalva Order Diptera Gastropoda Neanthes arenaceodentata Insecta Taxa studied Phylum MOLLUSCA Phylum ANNELIDA Class Insecta Phylum ARTHROPODA Table 1.8: The impacts of Literature review 31 1981 urger 1984 deen 1980 l. 1989 l. 1985 Mulla et al. 1982 Ali 1981 Reference Jackson et al. 1994, 2002 c 8 w Painter et al. 1996 Duration b C 3-28 d Ali 1981 OD 4 d Liber et al. 1998 EFC 4 m Hershey et al. 1995 EFC 1 d Purcell 1981 EFC 1 d Purcell 1981 EFC 3-28 d Ali 1981 EFC 3 d Dickman 2000 EFC 2 m Miura et al. 1980 EFC 3d Reyetal.1998 EFC 2-14 d Mulla et al. 1982 Dosage a F (B) EFC 4 d Car and de Moor 1984 F (M) USA F (B) EFC 4 d Molloy 1992 Canada F(B) EFC 7d ColboandUn USA F (B) EFC 2 d Pistrang and B USA F (B) EFC 6 d Merritt et a Canada F (B) EFC 3 d Back et a Hong Kong L EFC 3 dCanada Dickman 2000 M EFC 5d SebastienandBrust Bactimos WP USA M EFC 2-14 d VectoBac G USA F (M) Wettable powder USA M EFC 3-28 d Teknar VectoBac 12AS USA F (B) EFC 2 d Wettable powder USA F (M) VectoBac WP Comm. formulation South Africa Effect Formulation Location Site No impact Wettable powder USA L EFC No impact Wettable powder USA M EF No impact Liquid formulation USA M No impact Bactimos No impact Bactimos WP USA M 71% mortality VectoBac G USA M 75% mortality Bactimos WP France L Increased drift Teknar 100% mortality VectoBac 12AS Hong Kong L 18-88% mortality Wettable powder USA M sp. Erythemis simplicicollis Chironomidae Coleoptera Callibaetis Chironomus annularius Ephemeroptera Odonata Order Coleoptera Order Odonata Order Ephemeroptera Taxa studied Table 1.8 cont.

t 1981 Jackson et al. 2002 Reference Gharib and Hilsenhoff 1988 c 8 d Ali 1981 /S Garcia et al. 1980 5 d Gunasekaran et al. 2004 5 d 5 d 2 d Jackson et al. 2002 2 d Jackson et al. 2002 Duration b OD 5 d Gharib and Hilsenhoff 1988 OD OD EFC 1 d Purcell 1981 EFC 1 d Purcell 1981 EFCEFC 4 m Hershey et al. 1 1995 m Barnes and Chapman 1998 EFC 7d ColboandUndeen1980 EFC 6 d Lawler et al. 1999 EFC 3 d Dickman 2000 Dosage a L L USA USA Canada M EFC 5 d Sebastien and Brus USA F (B) EFC 6 d Merritt et al. 1989 VectoBac 12AS USA F (B) EFC 2 d Teknar HP-D USA L EFC Sandoz liquid Sandoz liquid Exp. formulation USA M EFC N Wettable powder USA M EFC 3-2 VectoBac 12AS USA F (B) EFC VectoBac 12AS USA F (B) EFC Effect Formulation Location Site No impact Comm. formulation Canada F (B) No impact No impact No impact VectoBac 12AS Australia F (M) No impact VectoBac G USA F (M) No impact Sandoz liquid USA L No impact Bactimos No impact VectoBac 12AS Hong Kong L No impact Wettable powder USA F (M) No impact Teknar No impact Wettable powder USA F (M) No impact VectoBac G USA F (M) sp. Elmidae Haliplidae Dytiscidae Hyallela azteca Notonecta Corixidae Araneae Hemiptera Gammaridae Talitridae Collembola Order Amphipoda Order Hemiptera Order Araneae Taxa studied Class Collembola Class Crustacea Crustacea Class Arachnida Table 1.8 cont. Literature review 33 Roberts 1995 Garcia et al. 1980 Reference c N/S Garcia et al. 1980 1-7 d 1-7 d Duration b OD 1d Brownetal.2000a OD 4d Brownetal.1996 OD 1 d Marten et al. 1993 EFC 3 y Niemi et al. 1999 EFC EFC EFC 3 d Dickman 2000 EFC 3 yEFC Niemi et al. 1999 2 m Miura et al.EFC 1980 3 y Niemi et al. 1999 EFC 1d Brownetal.1999 EFC 3 d Rey et al. 1998 EFC 2 m Miura et al. 1980 EFC 2 m Miura et al. 1980 EFC N/S Garcia et al. 1980 EFC 2 m Miura et al. 1980 Dosage ) a L L England England USA L OD 1 d Milan et al. 2000 USA L OD 2 d Bactimos powder Exp. formulation USA M EFC N/S Bactimos powder VectoBac G USA F (M) Effect Formulation Location Site No impact VectoBac G USA F (M) No impact Exp. formulation USA M EFC No impact Liquid formulation USA M No impact VectoBac 12AS Hong Kong L No impact VectoBac 12AS Australia F (M No impact No impact No impact Bactimos WP France L No impact Liquid formulation USA M No impact Liquid formulation USA M No impact VectoBac G USA F (M) No impact Exp. formulation USA M No impact Liquid formulation USA M 95% mortality VectoBac 12AS Australia L 50% mortality VectoBac 12AS Australia L 50% mortality Granule 50% mortality Granule 50-95% mortality VectoBac 12AS USA L sp. sp. sp. Copepods Hemigrapsus Neocardina serrata Clam shrimp Caradina indistincta Palaemonetes varians Leander tenuicornis Cyclops vernalis Simocephalus vetulus Daphnia magna Cyprois Daphnia pulex Macrocyclops albidus Macrocyclops Cladoceran Gammarus duebeni Ostracods Subclass Copepoda Order Decapoda Order Ostracoda Order Branchiopoda Taxa studied Table 1.8 cont.

2 Reference Garcia et al. 1980 verdosing concentration c rvae. Duration b FC 5 d Gunasekaran et al. 2004 OD 3 d Brown et al. 1998a OD 4 d Lee and Scott 1989 OD 1 d Snarski 1990 EFCEFC 3 y 3 y Hanowski et al. 1997 Niemi et al. 1999 Dosage a USA L EFC 2 d Tietze et al. 1991 Australia L OD 1 d Brown et al. 200 Canada L OD 3d Fortinetal.1986 uliidae (black fly larvae) that are considered as target species. . ration reached during operational procedures (Table 1.3); OD = O Exp. formulation USA M EFC N/S Bactimos itat of black fly larvae; (M) = Intertidal wetland habitat of mosquito la Effect Formulation Location Site No impact Teknar HP-D India L E No impact VectoBac G USA F (M) No impact Teknar No impact VectoBac G USA F (M) 50% mortality VectoBac 12AS Australia L 95% mortality VectoBac EC USA L 100% mortality VectoBac G USA L 20-86% mortality Teknar (fry) Birds Pseudomugil signifer Melanotaenia duboulayi Salvelinus fontinalis Pimephales promelas Agelaius phoeniceus Gambusia affinis Fundulus heteroclitus EFC = Concentrations equivalent to estimated field concent (5 to 1,000 times the estimated field concentration). Taxa studied Phylum CHORDATA Class Osteichthyes Class Aves Table 1.8 cont. d. Includes all diptera other than Culicidae (mosquitoes) and Sum b. c. d = days; w = weeks; m = months; y = years; N/S = not stated Notes a. L = laboratory; M = mesocosm; F = field; (B) = River hab Literature review 35

(Pistrang and Burger 1984). A moderate increase in the drift of Ephemeroptera was recorded at 24 h but not at 48 h by Pistrang and Burger (1984). The experiment incorporated a before/after design and drift samples were collected from three stations downstream. However, the drift patterns were poorly sampled prior to the treatment – only once, 24 h beforehand – additionally no untreated control data was collected. Thus, it is hard to state if the increase in drift was abnormal. No field trials have been published assessing the impact of Bti on Ephemeroptera in saltmarshes and mangroves.

Bti has not impacted on the true bugs (Hemiptera) in the majority of cases. In particular, waterboatman (Corixidae) were not affected in the laboratory (Dickman 2000), mesocosms (Ali 1981) or in flowing rivers (Jackson et al. 2002). Additionally, backswimmers (Notonectidae) were not affected in mesocosms (Ali 1981); however, a decrease in abundance in saltmarshes has been observed (Purcell 1981). This latter study used a before/after design; however, samples were only collected once before and after treatment, greatly reducing experimental strength as temporal variation was not accounted for.

The dragonflies and damselflies (Odonata) were not impacted in either the laboratory (Erythemis simpliciollis: Painter et al. 1996), mesocosms (Libellula, Anax and Gomphus: Sebastien and Brust 1981) or in saltmarshes in the USA (Anax sp. and Tramea sp.: Purcell 1981). Similarly beetles (Coleoptera) were not affected in the laboratory (Dytiscidae: Gharib and Hilsenhoff 1988, Haliplidae: Jackson et al. 2002), mesocosms (Ali 1981, Hydrophilidae and Dytiscidae: Mulla et al. 1982), in saltmarshes (Acilius sp. and Thermonectus sp.: Purcell 1981) and in lentic freshwater in the USA (Carabidae and Dytiscidae: Hershey et al. 1995).

Crustacea appear to be unaffected by Bti exposure at operational concentrations, and in Australia no effects were recorded in saltmarshes (Barnes and Chapman 1998).

No mortality of amphipods (Hyallela azteca) collected from lentic freshwater in the USA was recorded when they were exposed to Bti in the laboratory (Gharib and Hilsenhoff 1988) or in mesocosms (Garcia et al. 1980). Additionally in field trials in the USA, no effect of Bti was recorded on the family Talitridae in saltmarshes (Lawler et al. 1999) or the family Gammaridae in flowing rivers (Merritt et al. 1989, Jackson et al. 2002).

Laboratory trials simulating operational concentrations have reported no impact on prawns (Gammarus duebeni and Palaemonetes varians: Roberts 1995) and freshwater shrimp (Neocardina serrata: Dickman 2000). However, in Australia, the mortality of a saltmarsh (Leander tenuicornis) and a freshwater shrimp (Caradina indistincta) was recorded when exposed to concentrations of Bti at least 100 times greater than would be expected in the field (Brown et al. 1996, 1999, 2000a). This was most likely due to a reduction in water quality, as Roberts (1995) has demonstrated that Bti passed through the gut of shrimp unchanged and that defecated pellets were toxic to mosquito larvae the day after exposure.

In lentic freshwater in the USA, Bti was non-toxic to Copepoda (Hershey et al. 1998). Additionally, in mesocosms Macrocyclops sp. and Cyclops sp. were unaffected at operational concentrations of Bti (Garcia et al. 1980, Miura et al. 1980). In the laboratory, Macrocyclops sp. was impacted only when dosed at over 1,000 times the estimated field concentration of Bti (Marten et al. 1993). During this trial it was recorded that the copepods “were only harmed when the Bti was so thick that their movement was impaired”.

Similarly, mortality of waterfleas (Daphia sp.; Branchiopoda) was only recorded when exposed to an overdose in the laboratory (Milan et al. 2000). At operational concentrations, no increase in mortality was recorded in the laboratory (Rey et al. 1998), mesocosms (Garcia et al. 1980, Miura et al. 1980) or in lentic freshwater (Niemi et al. 1999).

Similar overdosing studies have been conducted in the laboratory with fish. The mortality of fish was only recorded when exposed to concentrations over 450 times that estimated to be reached in the field (Fortin et al. 1986, Lee and Scott 1989, Snarski 1990, Brown et al. 1998a). At such high concentrations, mortality can be attributed to depletion of dissolved oxygen rather than Bti exposure (Fortin et al. 1986, Snarski 1990). Additionally, xylene, used in formulations of Teknar to improve shelf life, affected the fish, rather than the Bti itself (Fortin et al. 1986). Literature review 37

1.7.2 s-methoprene

Much literature has been published about the impact of s-methoprene on non-target organisms (Glare and O'Callaghan 1998a, WHO 2000, Antunes-Kenyon and Kennedy 2001, Stark 2005a) and the possible impacts on macroscopic organisms have been summarised in Table 1.9. The studies were conducted in different environments: 58% (n = 30) in the laboratory, 31% (n = 16) in the field, and 11% (n = 6) in mesocosms. The impacts of normal operations were assessed by 58% of the studies, and the rest (42%) were assessed as an overdose.

In general, s-methoprene does not appear to impact on Mollusca. Gastropods (Physa sp. and Lymnaea sp.) were not impacted at operational concentrations in mesocosms (Schaefer et al. 1974), or when overdosed in the laboratory (Miura and Takashi 1973) and mesocosms (Creekmur et al. 1981). Additionally, neither Gastropods nor Bivalves were impacted in lentic freshwater in the USA (Hershey et al. 1995, 1998). However, the density of Gastropods (Physa sp.) was recorded to be reduced in the field by Breaud et al. (1977). As this trial lacked adequate spatial replication (utilising one treatment and control wetland), it is difficult to state if the results were abnormal.

S-methoprene has been recorded to impact on Dipteran species, particularly the family Chironomidae, as well as families Ephydridae, Psychodidae and Syrphidae (Miura and Takahashi 1973, Farghal and Temerak 1981). Regarding Chironomidae, s-methoprene was originally observed to inhibit the emergence of pupae during a laboratory trial by Miura and Takahashi (1973). This initiated research activity into the potential of s-methoprene for the control of pest chironomid species. Subsequently, s-methoprene was found to inhibit the emergence of Chironomus sp. and Tanypus sp. pupae in the laboratory (Mulla et al. 1974) and various chironomid species in mesocosms (Schaefer et al. 1974, Mulla et al. 1974, Norland and Mulla 1975). In mesocosms, liquid s-methoprene controlled Tanytarsini and Chironomus sp. for 2 weeks, and the solid formulations (pellets and briquets) provided extended control for 5 to 7 weeks (Ali 1991). Additionally, higher dosages of solid methoprene (Altosid pellets) provide more effective and extended control of Chironomus, Geoldichironomus and Tanypus spp. (Farghal and Temerak 1981).

0 77 . 1977 1985 d Mulla 1975 Ali 1991 Mulla et al. 1974 Creekmur et al. 1981 Reference c y Hershey et al. 1998 /S Schaefer et al. 1974 3 y Hershey et al. 1998 7 w 7 w 14 d 14 d Duration b FC 4 m Hershey et al. 1995 FC 3 y Hershey et al. 1998 OD 3 d Miura and Takahashi 1973 EFC EFC EFC EFC EFC 4 m Hershey et al. 1995 Dosage a F F F F F EFC 3 y Niemi et al. 1999 F EFC 9 d USA USA USA USA USA F EFC 18 m Breaud et al USA F EFC 18 m Breaud et al. 19 USA L OD 4 d Reish et al. USA F EFC 3 m Pinkney et al. 200 USA F EFC 3 m Norland an ALL Altosid pellets Altosid granules Altosid XR briquets Altodis 515225 USA M EFC N Altosid granules USA F EFC 3 Altosid powder USA M OD 2 w Altosid SR-10 USA M EFC 2 w ALL e Effect Formulation Location Site No impact Altosid briquets USA F E No impact Altosid ZR-515 USA L No impact Altosid No impact Altosid briquets USA F No impact Altosid granules USA F EFC 61-87% EI 38-98% EI 64-99% EI 30-100% EI Significant EI 50% reduction Altosid EC Reduced density Altosid Reduced density Altosid Reduced density Altosid granules USA F E -methoprene on non-target organisms. s spp. d Lymnaea & Chironomidae Bivalva Physa Physa Gastropoda Insecta Neanthes arenaceodentata Order Diptera Taxa studied Phylum MOLLUSCA Class Insecta Phylum ANNILIDA Phylum ARTHROPODA Table 1.9: The impacts of Literature review 39 1975 al. 2000 Floore et al. 1988 Creekmur et al. 1981 Hershey et al. 1995 Norland and Mulla 1975 Schaefer et al. 1974 Reference c d Majori et al. 1977 5 d 2 d Miura and Takahashi 1973 10 d Duration b C 3 d Farghal and Temerak 1981 FC 3 d Farghal and Temerak 1981 FC N/S Schaefer et al. 1974 OD 2 w Creekmur et al. 1981 OD 3 d Miura and Takahashi 1973 EFC 2 m Miura and Takahashi 1974 EFC N/S Schaefer et al. 1974 EFC EFC EFC 12 d EFC 12 d Miura and Takahashi 1973 EFC N/S EFC 12 d Miura and Takahashi 1973 EFC 3 w Lothrop and Mulla 1998 EFC 7 d Mulla et al. 1974 Dosage a L F USA F EFC 3m NorlandandMulla USA F EFC 3m Pinkneyet USA USA Altosid pellets USA M EFC 10 d Altosid pellets USA M OD 2 w Altosid briquets USA F EFC 4 m Altosid SR-10 Italy F EFC 14 Altosid EC Altosid 515225 USA M EFC N/S Altosid EC Effect Formulation Location Site 75% EI Altosid SR-10 Egypt F EF No impact Altosid SR-10 USA M No impact Altosid pellets USA M No impact Altodis SR-10 Egypt F E No impact Altosid EC No impact Altosid 515225 USA M No impact ALL Noimpact AltosidZR-515 USA L OD 50% mort. Altosid ZR-515 USA L 50% mort. Altosid ZR-515 USA L 60-100% EI Altosid ZR-515 USA L Significant EI Altosid 515225 USA M Significant EI Altosid 515225 USA M E 50% mortality Altosid ZR-515 USA L 50% mortality Altosid ZR-515 USA L >90% mortality Altosid pellets USA F 6-90% mortality 5-60% mortality sp.) sp. sp. sp. sp. sp. & Eristalis sp. sp. Notonecta unifasciata Hemiptera Dytiscus Helophorus Laccophilus Laccophilus Coleoptera Ephemeroptera Callibaetis pacificus Psycodidae Callibaetis Ephydridae Syrphidae ( Chironomus Geoldichironomus Chironomus stigmaterus Chironomidae Order Hemiptera Order Coleoptera Order Ephemeroptera Taxa studied Table 1.9 cont.

ney 1994 cKenney 1994 00a 999 Mulla 1975 1999 Forward and Costlow 1978 Reference c Duration b FC 10 w Lawler et al. 2000 FC 7 d Batzer and Sjogren 1986 OD 1 d Miura and Takahashi 1973 OD 4 d Gradoni et al. 1976 EFC 3 m Miura et al. 1978 EFC 3 m Miura et al. 1978 EFC 3 m Hershey et al. 1995 EFC 10 d Floore et al. 1988 Dosage a L OD N/S Christiansen et al. 1977 F EFC 4 d USA L OD N/S Celestial and M USA L EFC N/S Celestial and McKen USA F EFC 3 m Pinkney et al. 2000 USA L OD 4 d Reish et al. 1985 USA F EFC 3 m Norland and USA FUSA EFC F 3 m EFC Pinkney et al. 2000 3 d Lawler et al. Australia L OD 4 d Brown et al. 1996 USA F EFC 18 m Breaud et al. 1977 Australia L EFC 1 d Brown et al. 1 Australia L OD 1 d Brown et al. 20 Altosid ZR-515 USA L (C) EFC N/S ALL Effect Formulation Location Site No impact Altosid briquets USA M No impact ALL No impact Altosid briquets USA M No impact Altosid pellets USA F E No impact Altosid briquets USA F No impact Altosid SR-10 USA F E No impact ALL No impact Altosid EC No impact ALL No impact Altosid pellets USA M No impact ALL 50% mortality Altosid ZR-515 USA L 20% mortality Altosid 50% mortality ALL 95% mortality ALL Reduced density Altosid 50-90% mortality Technical Altosid Italy L (zoel) Mortality AL (zoel) 16-100% mortality Altosid ZR-515 USA Notonecta unifasciata Rhithropanopeus harrisii Buenoa scimitra Elasmopus bampo Eubranchipus bundyi Damselfly naiads Hyalella azteca Caradina indistincta Leander tenuicornis Trichocorixa reticulata Odonata Gammarus aequicauda Odonata naiads Talitridae Collembola Order Odonata Order Amphipoda Order Decapoda Taxa studied Class Collembola Class Crustacea Table 1.9 cont. Literature review 41 Matthews al. 1977 1990 Mortimer and Chapman 1995 Horst and Walker 1999 Miura and Takahashi 1973 McAlonan et al. 1976 Reference c 4 d 4 d 4 d 4 d Duration b C 3 y Niemi et al. 1999 D 2 d Bircher and Ruber 1988 OD OD OD 1 d OD OD EFC N/S Schaefer et al. 1974 EFC 14 d Majori et al. 1977 Dosage ) EFC 60 d Barber et al. 1978 a L L L L Australia Australia USA F EFC 18 m USA F EFC 18 m Breaud et USA L(C) OD 3d USA L OD 4 d Wirth et al. 2001 USA L OD 15 d McKenney and Australia Australia USA L OD 2 d Marten et al. 1993 ALL ALL N/S Tech. formulation USA L (C) EFC 11 d Altosid ZR-515 USA F EFC 5 d Altosid 10-F USA F EFC 10 w ALL ALL Effect Formulation Location Site formation No impact No impact Altosid 515225 USA M No impact N/S No impact Altosid SR-10 USA L (C No impact Altosid granules USA F EF No impact Altosid SR-10 Italy F No impact No impact 50% mortality 50% mortality Altosid ZR-515 USA L Changes in shell Reduced density Altosid Reduced density Altosid 40-74% mortality Altosid SR-90 USA L O 50-95% mortality ALL Increased mortality ALL sp. (nauplii) & Diatomus & sp. (nauplii) australiense Macrobrachium Heloecius cordiformis Mictyris longicarpus Apocyclops spartinus Palaemonetes paladosus Cyclops Palaemonetes pugio Palaemonetes pugio Copepods Palaemonetes pugio Uca pugilator Callinectes sapidus Cyclops Trypaea australiensis Taphromysis louisianae Subclass Copepoda Taxa studied Table 1.9 cont.

lanc 2003 l. 2006 Mulla 1975 nd LeBlanc 2001 iemi et al. 1999 McKenney and Celestial 1996 Reference c d Templeton and Laufer 1983 1 d Miura and Takahashi 1973 4 d 15 d Duration b FC 3 y Niemi et al. 1999 OD 1 d OD 1 d OD EFC N/S Schaefer et al. 1974 EFC 5 d EFC 5 d Miura and Takahashi 1973 EFC Dosage a L OD 2d Chuetal.1997 L L (C) USA L (C) EFC N/S Olmstead and LeB USA L (C) EFC N/S Olmstead a Hong Kong USAUSA FUSA EFC 3 m Norland and Belgium L (C) OD 10 d Ghekiere et a Technical Altosid ZR-515Technical USA L (C) OD 8 Altosid granules USA F EFC 3 y N ALL ALL Effect Formulation Location Site oocytes No impact Altosid 515225 USA M No impact Altosid granules USA F E No impact Altosid ZR-515 USA F No impact Altosid EC No Impact Altosid ZR-515 USA F 50% mortality Altosid ZR-515 USA L OD 50% mortality Altosid ZR-515 USA L 50% mortality Altosid ZR-515 USA L 50% mortality ALL Altered sex of 100% mortality Delayed molting Technical Reduced weight Reduced growth Reduced growth and no. of young and reproduction and reproduction sp. sp. Cladocera Daphnia magna Eulimnadia Moina macrocopa Cyprinotus Neomysis integer Mysidopsis bahia Triops longicaudatus Ostracods Order Cladocera Order Branchiopoda Order Ostracoda Order Mysidacea Taxa studied Table 1.9 cont. Literature review 43 89 1 02 l. 1998a Ellgaard et al. 1979 Miura and Takahashi 1974 Reference c verdosing concentration (5 to 3 y Niemi et al. 1999 2 w Miura and Takahashi 1973 37 d Ross et al. 1994b ecies. Duration b OD 1 d Miura and Takahashi 1973 EFC 10 w McAlonan et al. 1976 Dosage a Australia L OD 3 d Brown et a Australia L OD 1 d Brown et al. 20 USA L (C) EFC 3 d La Clair et al. 1998 USA L OD 4 d Lee and Scott 19 USA L EFC 2 d Tietze et al. 199 -methoprene metabolites Technical product USA L (C) OD s Altosid SR-10 USA L (C) OD 12 d ALL Altosid SR-10 USA F EFC 2 m ntration reached during operational procedures (Table 1.6); OD = O ed. ) and Sumuliidae (black fly larvae) that are considered as target sp Not stated. Effect Formulation Location Site No impact Altosid granules USA F EFC No impact ALL No impact ALL No impact Altosid ZR-515 USA L No impact Altosid ZR-515 USA L OD No impact Altosid 10-F USA F and weight No impact on 50% mortality Altosid EC locomotor activity Reduction in length Caused deformities (fry) & Melanotaenia duboulayi Birds Pseudomugil signifer Pimephales promelas Bufo boreas halophilus Xenopus laevis Gambusia affinis Fundulus heteroclitus Gambusia affinis Carassius auratus 1,000 times the estimated field concentration). e. EI = Emergence inhibition. d. Includes all diptera other than Culicidae (mosquitoes c. d = days; w= weeks; m= months;y =years; N/S= Notes a. L = laboratory; F = field; (C) = chronic impacts were studi b. EFC = Concentrations equivalent to estimated field conce Class Aves Taxa studied Phylum CHORDATA Class Osteichthyes Class Amphibia Table 1.9 cont.

These primarily mesocosm-based studies focus on the use of s-methoprene as a control agent against nuisance Chironomidae; thus, little insight is gained into the toxicity of s- methoprene against non-target Chironomidae. In a long-term field trial, Hershey et al. (1998) and Niemi et al. (1999) did record reduced densities of non-target Chironomidae after one year. In contrast, two well-designed field trials conducted in lentic freshwater in the USA, recorded no impact of s-methoprene on non-target Chironomidae (Hershey et al. 1995, Pinkney et al. 2000). Thus, the impact of s-methoprene on non-target Chironomidae is inconclusive; but high dosages and repeated applications are likely to have a detrimental effect.

Extensive trials indicate that other insects (aside from Diptera) are unaffected by s- methoprene exposure. S-methoprene was non-toxic to Ephemeroptera in the majority of cases. No mortality of Callibaetis sp. was observed when exposed to an overdose in the laboratory (Mirua and Takahashi 1973) or when exposed to operational concentrations in mesocosms (Schaefer et al. 1974) and lentic freshwater (Pinkney et al. 2000). One study did report an increased mortality of Ephemeroptera when exposed to relatively high dosages (0.1 and 0.05 ppm) during laboratory and mesocosms trials (Callibaetis pacificus: Norland and Mulla 1975). However, these mesocosm trials were not extensively replicated and only one pond served as a control.

Coleoptera and Hemiptera are not affected by s-methoprene exposure at operational concentrations. No mortality of a range of species has been recorded during mesocosm (Miura and Takahasi 1974, Schaefer et al. 1974, Miura et al. 1978, Creekmur et al. 1981) and field (Majori et al. 1977, Hershey et al. 1995, Norland and Mulla 1975, Lawler et al. 2000) trials in the northern hemisphere. Similarly, North American Odonata and Collembola were not affected at operational concentrations in the field (Norland and Mulla 1975, Hershey et al. 1995, Pinkney et al. 2000). However, the mortality of Coleoptera and Hemiptera was observed when exposed to at least 1,000 times the estimated field concentration of technical methoprene in the laboratory (Miura and Takahashi 1973).

S-methoprene does not cause mortality of Crustacea at operational concentrations. During laboratory trials, decapod species such as Leander tenuicornis (Brown et al. 1999), Rhithropanopeus harrisii (Christiansen et al. 1977, Celestial and McKenney 1994), and Palaemonetes pugio (McKenney and Matthews 1990), were unaffected when exposed to Literature review 45 operational concentrations, however, mortality was recorded when exposed to an overdose of at least 6 times the estimated field concentration. During field trials, no mortality of Leander tenuicornis (Brown et al. 1999), Eubranchipus bundyi (Batzer and Sjogren 1986) and Palaemonetes pugio (McAlonan et al. 1976) was recorded when exposed to operational concentrations of s-methoprene. However, the density of Palaemonetes paladosus and Taphromysis louisianea was reduced when exposed to operational concentrations in a poorly replicated field trial (Breaud et al. 1977).

Similarly, increased mortality was only recorded when Amphipoda, Copepoda, Cladocera, Branchiopoda and Mysidacea, were exposed to an overdose at least 6 times the estimated field concentration in the laboratory (Miura and Takahashi 1973, Gradoni et al. 1976, Reish et al. 1985, Bircher and Ruber 1988, Marten et al. 1993, Chu et al. 1997). At operational concentrations, no mortality of Amphipoda, Copepoda, Cladocera, Branchipoda and Ostracoda was recorded in mesocosms, lentic freshwater and saltmarshes (Schaefer et al. 1974, Miura and Takahashi 1973, Majori et al. 1977, Lawler et al. 1999, Niemi et al. 1999).

In addition, fish were unaffected when exposed to operational concentrations of s- methoprene in the laboratory (Miura and Takahashi 1973, Tietze et al. 1991) or field (Miura and Takahashi 1974, McAlonan et al. 1976), but were affected when overdosed, at least 10 times the estimated field concentration, in the laboratory (Lee and Scott 1989, Brown et al. 1998a, Brown et al. 2002).

The above results provide information on the acute effects of s-methoprene on non-target organisms, but disregard any chronic effects. The action of s-methoprene as a mosquito control agent depends on the hormone mimic interfering with normal neuroendocrine functions, and often takes up to 1 week for impacts to be noted on target organisms. Considering this, s-methoprene has potential to affect other organisms that undergo similar moult cycles to Diptera. However, many non-target impact studies have been conducted over a relatively short time, such as 1 to 4 days (e.g. Miura and Takahashi 1973, Farghal and Temerak 1981, Reish et al. 1985, Brown et al. 2000a) and this may not be sufficient time to record subtle effects.

During the 1970s, no chronic impacts on decapods or fish at estimated field concentrations were recorded (Barber et al. 1978, Forward and Costlow 1978, Ellgaard et al. 1979). More

46 recently, field concentrations have been reported to cause chronic impacts, including weight loss and delayed release of brood of the saltmarsh mysid, Mysidopsis bahia (McKenney and Celestial 1996), altered shell formation of the blue crab, Callinectes sapidus (Horst and Walker 1999), reduced growth and altered reproduction of the Branchiopoda, Daphnia magna (Olmstead and LeBlanc 2001, 2003; Templeton and Laufer 1983) and reduced length and weight of flathead minnow, Pimephales promelas (Ross et al. 1994b). These results suggest that s-methoprene may have chronic impacts on non-target organisms in the field.

There has been much controversy surrounding the impact of s-methoprene on frogs. This began after an outbreak of deformities in California, where s-methoprene had been regularly used for mosquito control. La Clair et al. (1998) publicised their belief that the by-products of s-methoprene metabolism were teratogenic and interfered with normal amphibian development. This claim was rebutted by Sullivan (1998), who indicated that the retinoic compounds formed by La Clair with a mercury lamp are unlikely to form in the field. He suggested that the outbreak of deformities was the result of a parasitic flatworm as recorded by Sessions and Ruth (1990).

1.8 Conclusions

Control of mosquito-borne disease is a challenge and global-warming will make the situation even more difficult. Therefore, there is an ongoing need for research to ensure that insecticides are used appropriately and efficaciously. Already a large body of literature exists which examines the efficacy of Bti, Bs, s-methoprene and pyriproxyfen. Yet there are gaps of knowledge surrounding the use of specific formulations in specific habitats: for example, VectoBac WG (Bti) and pyriproxyfen against the range of Australian mosquito larvae, VectoBac 12AS (Bti) in freshwater ground-pools, VectoBac G (Bti) in heavily vegetated areas and containers containing freshwater, or Altosid pellets (s-methoprene) in ground-pool freshwater habitats. Literature review 47

In general, effective control of freshwater larvae (ground-pool and container) has been intermittent, even though the adults are prominent vectors of pathogens. Regarding saltmarshes and mangroves, mosquito control is well developed and broad-scale control is carried out in Queensland, Western Australia and the Northern Territory, but treatment of heavily vegetated areas still poses challenges. The first focus of the thesis will be to undertake specific research to enhance the use of insecticides for mosquito control.

The non-target impacts of Bti and s-methoprene applications in Australian saltmarshes and mangroves remain to some extent unclear. It is important to note that much of the previous research has been based in the laboratory. While providing a great deal of information about the specific responses of different taxa to Bti and/or s-methoprene, these studies fail to account for ecological processes that may alter the effects of the pollutant, such as individual stress, population-level consequences or even the combination of other pollutants in the ecosystem (Underwood 1995, Chapman 2002). Also, studies conducted in mesocosms or with inadequate replication over a short time, do not account for natural population fluxes and may provide an unrealistic indication of non-target impacts.

Although limited Australian data exists, most assessments have been conducted in the northern hemisphere where environmental conditions may be different. It is inappropriate for managers to generalise this information for Australian saltmarshes, as the northern hemisphere saltmarshes are fundamentally different to those of Australia (Adam 2002). Given the broad-scale nature of Bti and s-methoprene insecticide use, it is imperative that that the ecological implications of management operations are understood; this will be the second focus of the thesis.

Chapter 2

Laboratory susceptibility of mosquito larvae to new biologically-based insecticides1

2.1 Introduction

For effective mosquito control, it is important that appropriate dosages are used in accordance with the susceptibility of the target larvae. The susceptibility of the target larvae is most easily defined in the laboratory. The methodologies used to assess the laboratory susceptibility of mosquito larvae to insecticides have been standardised by the World Health Organisation (see WHO 1996, 2005) and most studies follow these protocols (e.g. Skovmand and Sanogo 1999, Brown et al. 2001, Fillinger et al. 2003). Experiments conducted in the laboratory are relatively inexpensive and simple to conduct compared to field trials, but they minimise environmental influences that may affect product performance. Laboratory trials have been previously conducted for Bs and s-methoprene against Australian mosquito larvae (see Tables 1.4 and 1.6).

Regarding Bti, the liquid formulation has been tested in the laboratory and was highly effective against a range of species (see Table 1.3). The granular formulation of Bti (VectoBac G) is not amenable to laboratory trials due to the large size (3 – 8 mm dia.) of the particles and the commensurately large size of the vessels needed for laboratory tests. The newly registered water dispersible granule formulation of Bti (VectoBac WG) is designed to be applied as a liquid after the granules are mixed with water on-site. Preliminary trials found VectoBac WG to be effective against Cx. annulirostris in the laboratory (Muller et al.

1 Aspects of this chapter, regarding the laboratory susceptibility of mosquito larvae to VectoBac WG, have been published in the Journal of Economic Entomology 96: 1786-1791 (Appendix A). 50

2001) and the concentration resulting in 95% mortality was 102 Bti ITU/L. However, the susceptibility of other target species needs to be determined.

The susceptibility of Australian larvae to pyriproxyfen has not previously been determined. In the northern hemisphere, pyriproxyfen has been recorded to be effective at inhibiting the emergence of Aedes and Culex species (see Table 1.7).

The susceptibility of Australian mosquito larvae to the new Bti-based VectoBac WG and technical grade pyriproxyfen were assessed during the following chapter. It is necessary that each new formulation (e.g. VectoBac WG) is tested against the common target larvae, as specific formulation characteristics may affect activity.

In response to the aforementioned concerns, the information derived from these laboratory trials can be used to design subsequent field trials. The efficacy of a product may alter in the field where it is influenced by the environment and climate (Ramoska et al. 1982, Mulla et al. 1990, Becker et al. 1992). Understanding the influence of the environment on product activity provides base-line information for developing sound field trials. In response, the change in Bti efficacy resulting from exposure to sunlight and sediment was investigated.

Laboratory susceptibility 51

2.2 Methods

2.2.1 Laboratory bioassays

2.2.1.1 Mosquito test species

For both Bti and pyriproxyfen, toxicity studies were conducted on laboratory reared larvae. Specimens were derived from colonies maintained in the Queensland Institute of Medical Research (QIMR) insectary. The methods for maintaining the colonies were developed by Kay Marshall at QIMR and for Ae. notoscriptus have been described by Watson et al. (2000). The species tested were: • Ae. aegypti (colonised from Townsville in 2000), • Ae. notoscriptus (Brisbane 1995), • Ae. vigilax (Victoria Point 2000), • Cx. annulirostris (Brisbane 1997), • Cx. quinquefasciatus (Gold Coast 2001), and • Cx. sitiens (Coomera Island 1996).

The efficacy of Bti was also assessed against a second strain of Ae. aegypti (Torres Strait 2000).

The efficacy of pyriproxyfen was also tested against the F1 generation of Ae. vigilax (collected from Bribie Island 2006), Ae. aegypti (Cairns 2006) and Cx. quinquefasciatus (Brisbane 2006). Adults of Aedes vigilax were collected with carbon dioxide and 1-octen-3- ol (octenol) (Kemme et al. 1993) baited CDC-style light traps (Sudia and Chamberlain 1962). Aedes aegypti were collected as eggs that were deposited in ovitraps (dark coloured buckets containing aged tap water). Culex quinquefasciatus were collected as larvae from a polluted ground-pool. All specimens were transported to the QIMR insectary and Ae. aegypti and Cx. quinquefasciatus were reared through to adults. The adults were sorted to species, released into mesh cages and provided with 15% sucrose solution, fresh apple and human-blood via a membrane feeder. The eggs were collected from the cage and hatched for the bioassays. 52

2.2.1.2 General

The laboratory bioassays were based on the standard as described by the WHO (2005). Unless otherwise stated, each trial was conducted in five replicate glass beakers, each containing 20 third-instars in 200 ml of test concentration (Figure 2.1). Five beakers holding 20 larvae in 200 ml of water without product were used as controls. The salinity of the water was 0 parts per thousand (ppt) for the freshwater species (Ae. aegypti, Ae. notoscriptus, Cx. annulirostris and Cx. quinquefasciatus) and 10.5 ppt for the saltmarsh/mangrove species (Ae. vigilax and Cx. sitiens). The bioassays were conducted in tap water, which had been left to sit for at least 24 h. Saline water was made by diluting seawater (35 ppt) with aged tap water. Test specimens were individually removed from rearing trays via pipette and distributed randomly among the beakers. Water lost due to evaporation was replaced daily. The trials were maintained at 27°C with a relative humidity of 75% under a light:dark cycle of 14:10 h.

2.2.1.3 Bacillus thuringiensis var. israelensis

Concentration-response data were developed for VectoBac WG (Lot no. 63.878.PG, Valent Biosciences, Australia) against the six target species, detailed above. Initially, range-finding tests with a wide range of exposure concentrations (i.e. 0.002, 0.017, 0.171 ppm product) were conducted. Based on these tests, a narrow range of concentrations was then used to determine the lethal concentrations resulting in mortality rates of 50% and 95% (LC50 and

LC95, respectively).

To treat the replicate beakers, a stock solution of VectoBac WG was created; 20 mg of VectoBac WG was weighed using microbalance scales (Mettler Toledo AT261 DeltaRange) and placed into 1 L of distilled water in a glass beaker. This mixture was stirred vigorously for several minutes until all of the VectoBac WG had visibly dissolved into the mixture. Assuming that the mixture was homogenous, 1 ml of the stock solution would contain 0.02 mg of VectoBac WG equivalent to 60 Bti ITU (1 mg of product contains 3000 Bti ITU). Using these calculations, each of the replicate beakers was treated to create the desired concentration. Laboratory susceptibility 53

Figure 2.1: Static exposure bioassays were conducted in replicate glass beakers each containing 20 test larvae.

The mortality of the target larvae was assessed at 48 h post treatment. Death or the lack of reaction to gentle prodding with a glass pipette was the measured deleterious response. The test larvae were not fed during testing to minimise variation due to nutrition and metabolic condition.

2.2.1.4 Pyriproxyfen

Concentration-response data were developed for technical grade pyriproxyfen against the six target species, detailed above. Initially, larvae were exposed to a wide range of pyriproxyfen concentrations (i.e. 0.000049, 0.485 and 43.65 ppb AI). Based on these tests, a narrow range of concentrations was then used to determine the dosages where 50 and 95% emergence inhibition of pupae was observed (EI50 and EI95, respectively).

As technical grade pyriproxyfen has low solubility in water, stock solutions (485 ppm AI) were made by dissolving 10 mg of technical grade pyriproxyfen (Lot no. 50401, 97% pure, Sumitomo Chemical Co. Australia) in 20 ml of acetone (Merck Pty Ltd, Victoria, 99.5% pure). To add the same amount of acetone to test and control beakers, working dilutions were made of the pyriproxyfen-stock-solution in acetone (48.5 – 174.6 ppm AI); and the appropriate amounts of pyriproxyfen-working-solution was added to the beakers. To examine the effects of acetone on the larvae, the same amount of acetone was added to five beakers without pyriproxyfen.

Pilot trials were conducted according to methods described by Chism and Apperson (2003), and beakers contained 500 µl of acetone in 195.5 ml of water. However, significant mortality of the acetone control larvae was observed. Thus, the amount of acetone was reduced to 10 µl of acetone per beaker. No significant mortality of acetone control larvae was observed when exposed to these concentrations of acetone compared with untreated larvae (student’s t-test: p <0.05, low-dose acetone control and water only control).

Mortality, including dead larvae, dead pupae and failed emergence, was assessed every 24 h. Each beaker was individually covered with a fine cotton mesh to contain any emerged adults. The larvae were fed during the assays on a per larva basis. Aedes aegypti and Cx. sitiens were fed a mixture of 50% lactalbumin powder (Sigma Chemical, St Louis, MO, Laboratory susceptibility 55

USA) and 50% yeast powder mixed in distilled water. Culex annulirostris, Cx. quinquefasciatus, Ae. notoscriptus and Ae. vigilax were fed ground fish food pellets (Wardley Pond TEN Pellets; Secaucus, NJ, USA). The different food types were observed to provide optimal growth of the respective species.

When third-instar Cx. annulirostris were placed in 200 ml beakers, significant (>30%) mortality of the larvae was observed. As such, the Cx. annulirostris trial was conducted in 500 ml of aged tap water in white plastic containers (6 cm high x 17 cm long x 12 cm wide), and individually aerated. Twenty first-instars were placed into each container. Evaporated water was replaced daily until the larvae were treated with pyriproxyfen as third-instars. The remainder of the experiment was conducted as described above.

2.2.2 Environmental influences

2.2.2.1 Effect of sunlight

The effect of sunlight on the duration of Bti activity against Ae. notoscriptus larvae was examined. Larvae were obtained from a colony maintained in the QIMR insectary (colonised from Brisbane in 1995). The duration of Bti activity was assessed in the laboratory and in a non-shaded outdoor setting.

The trials were conducted in replicate terracotta pot plant saucers, 15 cm dia., containing 200 ml of tap water (left to sit for at least 24 hours). Pot plant saucers were chosen because in north Queensland they provide habitat for Ae. aegypti and Ae. notoscriptus. Four replicate assays were conducted for each treated and untreated control saucer. To assess the duration of control, 20 third-instars were added to each treated and untreated saucer on a weekly basis, and mortality was assessed after 48 h. At this point all surviving larvae and/or cadavers were removed. This procedure was repeated weekly until no significant difference in mortality was recorded between untreated control and treatments.

Larvae were exposed to 0.04 g VectoBac G/saucer; 6 µl VectoBac 12AS/saucer and 2.6 mg of VectoBac WG/saucer. The dosages equated to 36,000 – 40,000 Bti ITUs/L. These were high dosages, at least 4 times the label rate, but due to the size of the VectoBac G granules, 56 this was the smallest amount possible and equivalent amounts of the other products were used.

2.2.2.2 Effect of sediment

The effect of sediment on the duration of Bti activity against Ae. aegypti was examined. Larvae were obtained from a colony maintained at the QIMR insectary (colonised from Townsville in 2000).

Trials were conducted using plant pots and matching saucers. The pots, 15 cm dia. and slightly tapered at the bottom, were filled with potting mix (GardenLovers, Australia) to a height of 14 cm. VectoBac G was spread over the soil surface. Four replicate assays for each treatment were conducted, consisting of 0.1, 0.5 and 0.75 g VectoBac G/pot. High dosages of product were used, equating to 20,000, 100,000 and 150,000 Bti ITUs/pot, respectively.

Each trial consisted of weekly assays where 20 third-instars were exposed to leachate in saucers and the number of surviving larvae at 48 h was recorded. At this time all surviving larvae and/or cadavers were removed from the saucers. Throughout the trial, four replicate untreated pots and saucers were treated in the same manner to provide control data. Larvae were added to the saucers on a weekly basis, until no statistically significant difference in mortality was recorded between untreated control and treatments.

For positive control data, the mortality of Ae. aegypti exposed to VectoBac G placed directly into saucers without soil was assessed. The trials were conducted in the same manner and consisted of four replicate treatments, each 0.5 g VectoBac G/saucer, and four untreated controls.

2.2.3 Statistical methods

The effect of VectoBac WG and pyriproxyfen concentration on mortality was assessed using probit analysis (Finney 1971; PROC PROBIT, SAS Institute 2001). Log-linear mortality or emergence inhibition was plotted using either the Normal model or the Laboratory susceptibility 57

Gompertz model, to provide the best fitting curve and most reliable analysis (Spickett and Van Ark 1990). This method was more appropriate than standard probit analysis, as the data were not normally distributed. This procedure was adopted in favour of Abbott’s formula (Abbott 1925) because it did not modify the exposure variable, and thus had negligible impact on the curve.

Data on the effect of sunlight and sediment on mortality of larvae in the various treatments were compared using a univariate ANOVA. The square root of the proportion of mortality was arcsine transformed to normalise the data (Anscombe 1948). Where necessary, an all- pairwise comparison test (Tukey 1953, cited in Zar 1999) was used to determine the significant differences for each factor. To compare the differences of treatment efficacy, with and without sunlight, a non-parametric Kruskal-Wallis test was used. To create figures and present means, data were back-transformed (Anscombe 1948, Zar 1999).

2.3 Results

2.3.1 Laboratory bioassays

2.3.1.1 Bacillus thuringiensis var. israelensis

All of the species tested were highly sensitive to VectoBac WG. After 48 h exposure third- instars of the freshwater species, Cx. annulirostris and Cx. quinquefasciatus, were found to be the most sensitive, with LC50 values of 12 ITU/L (0.004 ppm) and 15 ITU/L (0.005 ppm), respectively, and identical LC95 values of 57 ITU/L (0.019 ppm) (Table 2.1, Figure

2.2). The saltmarsh species, Ae. vigilax, was slightly more tolerant with an LC50 value of 39

ITU/L (0.013 ppm) and LC95 values of 63 ITU/L (0.021 ppm). Culex sitiens, a co-habitant of saltmarshes, recorded the highest LC50 value of 57 ITU/L (0.019 ppm) and an LC95 value of 93 ITU/L (0.031 ppm). For the container breeding species, Ae. aegypti, LC50 and LC95 values of 54 ITU/L (0.018 ppm) and 78 ITU/L (0.026 ppm) were estimated, respectively for the Torres Strait strain and 51 ITU/L (0.017 ppm) and 93 ITU/L (0.031 ppm), respectively for the Townsville strain. Aedes notoscriptus had an LC50 value of 45 ITU/L (0.015 ppm) and the highest LC95 value of 111 ITU/L (0.037 ppm).

2.3.1.2 Pyriproxyfen

After exposure to pyriproxyfen in the laboratory, the colony strains of the freshwater species Cx. annulirostris and the saltmarsh species Cx. sitiens were both found to be highly sensitive, with EI50 values of 0.003 and 0.001 ppb AI, respectively, and EI95 values of 1.49 and 1.51 ppb AI, respectively (Table 2.2; Figure 2.3). The container breeding species were more tolerant to pyriproxyfen. The colony and F1 strains of Ae. aegypti recorded EI50 values of 0.28 and 0.92 ppb AI, respectively, and EI95 values of 17.19 and 17.51 ppb AI, respectively. The colony strain of Ae. notoscriptus recorded an EI50 value of 2.87 ppb AI and an EI95 value of 25.93 ppb AI. For the freshwater species, Cx. quinquefasciatus, the EI50 and EI95 values of the Gold Coast colony strain was 0.13 and 25.93 ppb AI, respectively, and for the F1 Brisbane specimens were 3.34 and 31.41 ppb AI, respectively. The Victoria

Point colony strain of Ae. vigilax had an EI50 value of 0.2 ppb AI and an EI95 value of 39.75 ppb AI and the F1 Bribie Island strain recorded an EI50 value of 1.34 ppb AI and an EI 95 value of 22.59 ppb AI. Laboratory susceptibility 59

c Slope ± SE b (df) 2 x al 199(58) 5.9±0.7 ormal 22 (57) 4.6 ± 0.4 Normal 132(53) 3.6±0.4 Distribution ) Gompertz 280(43) 6.5±0.7 – 0.036) Gompertz 158 (68) 2.3 ± 0.2 ITU/mg) after 48 h exposure. 19 – 0.024) Normal 113 (43) 6.5 ± 0.8 Bti 0.024 – 0.029) Gompertz 280 (98) 0.3 ± 0.03 (95% CL) 95 LC oBac WG (3,000 a (95% CL) 50 0.004 (0.003 – 0.005) 0.019 (0.015 – 0.036) N 0.015 (0.013 – 0.017) 0.037 (0.032 – 0.046) 0.017 (0.015 – 0.018) 0.031 (0.023 – 0.037) Norm Coomera Island 0.019 (0.018 – 0.021) 0.031 (0.023 – 0.036 Brisbane Gold Coast 0.005 (0.003 – 0.006) 0.019 (0.015 Victoria Point 0.013 (0.012 – 0.014) 0.021 (0.0 Brisbane ColonyoriginTownsville LC Torres Strait 0.018 (0.016 – 0.019) 0.026 ( b. df = degreesc. of freedom SE = standard error Notes a. Concentrations in ppm product; CL = confidence limits Cx. quinquefasciatus Cx. sitiens Cx. annulirostris Ae. vigilax Ae. notoscriptus Species Ae. aegypti Ae. aegypti Table 2.1: Laboratory susceptibility of mosquito larvae to Vect 60

1.00 Ae. aegypti Ae. aegypti (Townsville) (Torres Strait) 0.75

0.50

0.25

0 10 100 1000 1 10 100 100

1.00 Ae. notoscriptus Ae. vigilax (Brisbane) (Victoria Point) 0.75

0.50

0.25

0 1 10 100 1000 10 100 1000 instar mortality 1000 -

1.00 Cx. annulirostris Cx. quinquefasciatus (Brisbane) (Gold Coast) 0.75 Probability of third 0.50

0.25

0 1 10 100 1 10 100 1000 Dose (Bti ITU/L) 1.00 Cx. sitiens (Coomera Island) 0.75

0.50

0.25

0 1 10 100 1000 10000 Dose (Bti ITU/L)

Figure 2.2: Mean mortality (±95% CI) of third-instars exposed to VectoBac WG (3,000 Bti ITU/mg) for 48 h in the laboratory. Laboratory susceptibility 61

c 0.19 0.37 0.10 0.11 0.07 0.17 0.08 0.07 0.08 ± ± ± ± ± ± ± ± ± Slope ± SE 1.14 1.50 1.19 0.82 0.53 1.74 0.55 0.63 0.63 0) ) b (df) 2 x Normal 150.6 (37) Gompertz 130.03 (33) Gompertz 160.8 (35) Gompertz 90.1 (31) Distribution 10.78) (95% CL) 95 EI 17.51 (9.70 – 48.38) 22.59 (16.25 – 32.92) Gompertz31.41 (14.60 – 209.33) 61.78 (43 Gompertz 221.76 (3 17.19 (7.29 – 61.22) 19.76 (14.10 – 31.45) Gompertz 210.3 (63) 39.75 (22.15 – 88.21) Gompertz 176.12 (65) 25.92 (10.8 – 101.73) Gompertz 74.6 (36) nical grade pyriproxyfen. a (95% CL) 50 0.92 (0.37 – 1.59) 0.28 (0.10 – 0.60) 2.87 (2.23 – 3.61) 0.003 (0.0008 – 0.007) 1.49 (0.45 – 0.96) Cairns Bribie Island 1.34 (0.78 – 2.03) Brisbane 3.34 (1.89 – 6.23) 1 1 1 Colony origin EI Townsville F Brisbane Brisbane Coomera Island 0.001 (0.0002 – 0.005) 1.51 (0.44 – Victoria Point 0.02 (0.08 – 0.39) Gold Coast 0.13 (0.06 – 0.24) F F Table 2.2: Laboratory susceptibility of mosquito larvae to tech Notes a. Concentrations in ppbb. AI; CL - confidence df limit = degreesc. of freedom SE = standard error. Species Ae. aegypti Ae. aegypti Ae. notoscriptus Cx. sitiens Cx. annulirostris Cx. quinquefasciatus Ae. vigilax Ae. vigilax Cx. quinquefasciatus 62

1.00 Ae. aegypti Ae. aegypti (Townsville) (F1 Cairns) 0.75

0.50

0.25

0 0.0001 0.001 0.01 0.1 1 10 100 0.001 0.01 0.1 1 10 100

1.00 Ae. notoscriptus (Brisbane) 0.75

0.50

0.25

0 0.0 0.1 1 10 100 Probability of emergence inhibition

1.00 Ae. vigilax Ae. vigilax (Victoria Point) (F1 Bribie Island) 0.75

0.50

0.25

0 0.0001 0.01 1 100 0.001 0.01 0.1 1 10 100 1000 Dose (ppb AI) Dose (ppb AI)

Figure 2.3: Mean emergence inhibition (±95% CI) of mosquito larvae exposed to technical grade pyriproxyfen in the laboratory.

Laboratory susceptibility 63

1.00 Cx. annulirostris Cx. sitiens (Brisbane) (Coomera Island) 0.75

0.50

0.25

0 0.00001 0.0001 0.001 0.01 0.1 1 10 0.00001 0.001 0.01 0.1 1 10

1.00 Cx. quinquefasciatus Cx. quinquefasciatus (Gold Coast) (F1 Brisbane) 0.75 Probability of emergence inhibition 0.50

0.25

0 0.0001 0.001 0.01 0.1 1 10 0.001 0.01 0.1 1 10 100 Dose (ppb AI) Dose (ppb AI)

Figure 2.3 cont. 64

2.3.2 Environmental influences

2.3.2.1 Effect of sunlight

With sunlight absent, significant mortality of Ae. notoscriptus exposed to the three Bti treatments (VectoBac WG, 12AS and G) was recorded for 7 weeks (ANOVA: p <0.0001; post-hoc: p <0.05; Figure 2.4). During this time, the mean (± SD) mortality of the treated larvae ranged from 81.4 ± 5.8% to 100 ± 0%. At 8 weeks, the mortality of larvae exposed to

VectoBac G and 12AS remained different from the untreated controls (ANOVA: F3,16 = 14.14, p <0.0001): VectoBac G, mortality recorded at 64.2 ± 5.3% (post hoc: p = 0.003), and VectoBac 12AS, recorded at 95.3 ± 4.6% (post hoc: p <0.0001). At this point in time, the mortality of larvae exposed to VectoBac WG, recorded at 30.1 ± 32.1%, was not statistically different from the untreated controls (post hoc: p = 0.068). After 8 weeks, the products did not produce effective control of the target larvae and mortality ranged from 0.9 ± 4.2% to 43.9 ± 17.5%. Untreated control mortality ranged between 0 ± 0% and 18.5 ± 8.7% during the trial.

When third-instar Ae. notoscriptus were exposed to the VectoBac formulations in sunlight, the duration of activity was significantly reduced. Mortality of larvae exposed to the three treatments was only different from the untreated controls for 48 h (ANOVA: F3,16 = 342.99, p <0.0001; post hoc: p <0.0001). At 1 week, mortality of larvae exposed to VectoBac G, recorded at 94.0 ± 9.9%, remained statistically different from the untreated controls

(ANOVA: F3,16 = 33.94, p <0.0001; post hoc: p <0.0001) and was the most effective product over time (Figure 2.4). VectoBac 12AS and VectoBac WG did not control the larvae at this point in time, mortality being recorded at 1.8 ± 1.5% (post hoc: p = 0.997) and 16.7 ± 2.7% (post hoc: p = 0.341), respectively. Untreated control mortality ranged between 0.8 ± 1.5% and 17.9 ± 1.9% throughout the trial.

The mortality of larvae exposed to VectoBac WG and VectoBac 12AS in sunlight at 1 week was statistically different from that recorded in the laboratory (sunlight absent; Kruskal- Wallis: p <0.05). The residual control of VectoBac G was not different between sunlight exposures until 2 weeks (Kruskal-Wallis: p <0.05). Laboratory susceptibility 65

a) sunlight absent 100

Ae. notoscriptus 0 0123456789101112

b) sunlight present 100

80 Control 60 VectoBac G VectoBac WG 40 VectoBac 12AS Percent mortality of third-instar 20

0 0123

Time post treatment (weeks)

Figure 2.4: Mean (± SD) percent mortality of third-instar Aedes notoscriptus exposed to VectoBac products (G: 200 Bti ITU/mg; WG: 3,000 Bti ITU/mg; 12AS: 1,200 Bti ITU/mg); a) with and b) without the presence of sunlight. Arcsine data are back transformed.

2.3.2.2 Binding to sediment

The low dose of VectoBac G (0.1 g/pot) did not leach through the soil and there was no significant difference in mean (± SD) mortality of target larvae, recorded at 6.3 ± 3.7% compared to the untreated controls at 48 h (post hoc: p = 0.968).

The mortality of larvae exposed to leachate of the medium dose (0.5 g/pot) was recorded at 46.32 ± 16.7% after 48 h and was significantly higher than untreated controls (ANOVA:

F3,40 = 6.83, p = 0.001; post hoc: p <0.0001; Figure 2.5). Leachate of active ingredient from this dosage ceased at 1 week and mortality was recorded at 4.9 ± 20.0%.

Some of the active ingredient from the high dose (0.75 g/pot) did leach through the soil with time. Mortality of target larvae was recorded at 9.3 ± 24.9% at 48 h, which increased to 33.8 ± 40.9% by 2 weeks, and decreased to 5.5 ± 2.9% by 4 weeks. Due to the high disparity in the mortality of different replicates, there was no statistical difference in mortality when compared with the untreated controls (post hoc: p >0.05). However, these results indicate that at higher dosages some of the active ingredient does not leach through soil. Untreated control mortality ranged from 0.3 ± 1.3% to 3.1 ± 2.6% throughout the trial.

The mortality of Ae. aegypti was significantly higher when VectoBac G was placed directly into saucers at 48 h (ANOVA: F1,10 = 14.5; p = 0.003) and 1 week (ANOVA: F1,10 = 32.0; p <0.0001). Without the influence of soil, the duration of Bti activity was significantly longer, being 12 weeks (Figure 2.5). Laboratory susceptibility 67

a) soil absent 100 Control 80 0.5 g

20 Ae. aegypti 0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

b) soil present 100

80 Control 60 0.1 g 40 0.5 g Percent mortality of third-instar 0.75 g 20

0 01234

Time post treatment (weeks)

Figure 2.5: Mean (± SD) percent mortality of third-instar Aedes aegypti exposed to VectoBac G (200 Bti ITU/mg); a) with and b) without the presence of soil. Arcsine data are back transformed. 68

2.4 Discussion

Based on the results of these laboratory trials, the new Bti formulation, VectoBac WG, shows promise as an effective mosquito control agent for use in Australia. Other Bti-based formulations are currently used for mosquito control, however unlike VectoBac WG, the susceptibility of different species has been recorded to be highly variable. During the laboratory trial, all mosquito species tested were sensitive to VectoBac WG, with the most sensitive species (Cx. annulirostris and Cx. quinquefasciatus) being twice as susceptible as the most tolerant (Ae. notoscriptus), indicated by LC95 values.

In contrast, greater variation in susceptibility to VectoBac 12AS was recorded in the laboratory; where Ae. aegypti (LC95: 886.46 Bti ITU/L) was indicated to be at least 36 times more tolerant of Bti than Cx. sitiens (LC95: 24.37 Bti ITU/L) by Brown et al. (1998b, 2001). In terms of product efficacy, VectoBac WG was found to be more efficient than VectoBac 12AS, assessed by Brown et al. (2001), when applied against the container breeding species,

Ae. aegypti, Ae. notoscriptus (LC95: 421.07 Bti ITU/L) and the saltmarsh/mangrove species

Ae. vigilax (LC95: 110.84 Bti ITU/L). Such variation in product efficacy may be related to formulation characteristics (Brown et al. 2001, Fillinger et al. 2003). Based on these results, the Bti-based VectoBac WG appears to be an excellent choice for controlling the range of target larvae common in Australia.

The duration of Bti activity was drastically reduced in the presence of both sunlight and sediment. My results indicated that Bti activity was 48 h in sunlight, compared with 7 weeks without sunlight. A similar result was recorded by de Melo-Santos et al. (2001) in Brazil, where the residual activity of a Bti tablet was 20 days less in the presence of strong sunlight. Becker et al. (1992) and Nayar et al. (1999) have indicated that the amount of Bti required for control at 48 h needs to be at least 3 times higher in sunlight compared with shade.

During the sediment trials, the active ingredient of Bti bound strongly to soil particles and did not readily leach. Similarly, Bti activity was reduced in laboratory bioassays containing sediment by Ramoska et al. 1982, Sheeran and Fisher 1992 and Yousten et al. 1992. Interestingly, Boisvert and Boisvert (1999) found that Bti protoxins bound to soil particles, when fed to larvae, maintained their activity for up to 5 months. Laboratory susceptibility 69

Regarding pyriproxyfen, all species were susceptible to the technical grade product, but there was significant variation in the susceptibility of species. The EI95 values calculated for the two most susceptible species (Cx. sitiens: 1.51 ppb and Cx. annulirostris: 1.49 ppb) were similar to those reported in the published literature for Culex tarsalis (Coquillett), a congener from the USA (0.14 ppb AI: Estrada and Mulla 1986, 0.25 ppb AI: Schafer et al. 1988).

The most tolerant species to pyriproxyfen exposure were the colony strains of Ae. vigilax

(EI95: 39.75 ppb AI), Ae. aegypti (EI95: 17.19 ppb AI) and Cx. quinquefasciatus (EI95: 25.29 ppb AI). In contrast, the laboratory-based EI95 value recorded in the USA for Ae. aegypti was 2.6 ppb AI (Estrada and Mulla 1986) and for Cx. quinquefasciatus was 0.16 ppb AI (Schaefer et al. 1988, Mulligan and Schaefer 1990) when exposed to technical grade pyriproxyfen. The Australian strains of these species were at least 6 and 86 times more tolerant to pyriproxyfen exposure, respectively, than their North American counterparts. As different statistical techniques were used to calculate the EI95 values, it is conceivable that this may have influenced the results. However, during these trials, high adult emergence was recorded even after exposure to relatively high amounts of pyriproxyfen, indicating that these were real differences in susceptibility. For example, when exposed to 24.25 ppb AI pyriproxyfen, 25 – 40% of Ae. aegypti emerged and 5 – 15% of Ae. vigilax emerged. It is unknown why the Australian strains of these species were more tolerant to pyriproxyfen exposure.

Pyriproxyfen appears to be less active than the currently registered insect growth regulator, s-methoprene, against Australian mosquito larvae. Ritchie et al. (1997c) determined the laboratory efficacy of s-methoprene to colonies of mosquito species derived from similar regions of Australia. Aedes aegypti, Ae. notoscriptus, Ae. vigilax and Cx. annulirostris were all sensitive to s-methoprene exposure; EI90 values ranged from 0.34 to 1.3 ppb AI (Table

1.5). Culex sitiens was indicated to be the most tolerant species with an EI90 value of 6.54 ppb AI; a much lower value than recorded for Ae. notoscriptus, Ae. vigilax, Cx. quinquefasciatus and Ae. aegypti when exposed to pyriproxyfen in the current trial. In contrast, Nayar et al. (2002) found that pyriproxyfen was more active than s-methoprene against Ae. aegypti and Cx. quinquefasciatus in both the laboratory and field in the USA. 70

Various formulations of pyriproxyfen have been registered for use by control programs overseas at 10 – 100 g AI/ha (6.6 – 66.6 ppb AI in 15 cm deep water; Schaefer et al. 1991, Schaefer and Mulligan 1991). Based on our results, these rates would provide control of Cx. sitiens and Cx. annulirostris, as the EI95 values (1.49 and 1.51 ppb AI, respectively) are at least 4 times less than the lowest estimated field concentration. Suggesting that the product would be useful for control in ground-pool freshwater habitats, where Cx. annulirostris is the target species.

Considering that the major target species of the Australian mosquito control industry is Ae. vigilax, the susceptibility of this species is of particular interest. However, it was the least susceptible species to pyriproxyfen. At the rates used overseas, pyriproxyfen would not be effective against Ae. vigilax, as the EI95 values for the colony and F1 strains (39.71 and 22.59 ppb AI) were at least three times the lowest estimated field concentration. As Chapman et al. (1999) found Ae. vigilax to be panmictic, a uniform tolerance to pyriproxyfen in its range throughout Australia can be assumed. This should be verified, however, for different geographic populations.

Determining the laboratory susceptibility of the common target species has provided useful baseline information for the development of sequential field trials and control operations. The new Bti formulation, VectoBac WG, was effective against the full range of species tested. However, dilutions of technical grade pyriproxyfen were only effective against certain species (Cx. sitiens and Cx. annulirostris). Therefore, it is likely that of the two products tested only the Bti-based VectoBac WG will be adapted into broad scale mosquito control operations in Australia. Before this happens, it is essential to test the efficacy of VectoBac WG in the field, especially since the presence of sunlight and sediment were demonstrated to significantly reduce Bti activity. 71

Chapter 3

Efficacy of Bacillus thuringiensis var. israelensis to control mosquito larvae under field conditions1

3.1 Introduction

After the base-line susceptibility of mosquito larvae to an insecticide has been determined in the laboratory, the insecticide should be tested in the field before it is recommended for use in routine control programs. The results from Chapter 2 indicate that the water dispersible formulation of Bti (VectoBac WG) will be useful for broad-scale mosquito control; however, the results were not so promising for pyriproxyfen. Thus further field trials with pyriproxyfen are unnecessary and this chapter will assess the use of Bti in the field.

The liquid (VectoBac 12AS; 1,200 Bti ITU/mg) and water dispersible (VectoBac WG; 3,000 Bti ITU/mg) Bti formulations were recorded to be highly effective against a range of Australian mosquito larvae in the laboratory (Table 1.3; Chapter 2). During field-based studies in saltmarshes, VectoBac 12AS was reported to be effective against Ae. vigilax (Mottram et al. 1989, Brown et al. 2001, Webb and Russell 2001), Cx. sitiens (Brown et al. 1998b, Muller et al. 2001) and Ve. funerea (Jeffery et al. 2005) (see Table 1.3). Similarly, VectoBac WG was highly effective against Ae. vigilax and Cx. sitiens (Muller et al. 2001) in the field. However, the efficacy of these products in ground-pool freshwater habitats, where the primary target species is Cx. annulirostris, requires assessment.

1 Various aspects of this chapter, regarding the efficacy of Bacillus thuringiensis var. israelensis for the control of mosquito larvae in freshwater and saltmarsh ground-pools, have been published in the Journal of Economic Entomology 96: 1786-1791 and the Proceedings of the 9th Arbovirus Research in Australia Symposium, pp 337-343 (Appendix A). 72

The granular Bti formulation (VectoBac G; 200 Bti ITU/mg) is registered only for use in saltmarshes and mangroves, and has been recorded to control Ae. vigilax (Ritchie 1994) and Ve. funerea (Jeffery et al. 2005). This product is useful for application via helicopter where there is heavy vegetation as the granules bounce through the canopy. However, the low potency and high bulk density of VectoBac G means that large amounts of product must be carried to treat a small area, and this leads to more expensive application costs.

One possible way to overcome the bulk density problem associated with VectoBac G is to lower the label rate, thus reducing the amount required to treat an area. When the label rates of the different Bti (VectoBac) formulations are compared in terms of potency (Table 3.1), it is evident that the VectoBac G label rate (7 kg/ha; 933 Bti ITU/L in 15 cm deep water) equates to the higher label rates of VectoBac 12AS (1.2 L/ha; 1,036 ITU/L) and VectoBac WG (500 g/ha; 1,000 ITU/L). However, VectoBac 12AS and VectoBac WG are often applied at the lower label rates. This suggests that the label rate of VectoBac G could be reduced. In a preliminary trial, Ritchie (1994), recorded VectoBac G to be effective below the label rate (97.2% mortality at 666 Bti ITU/L; 5kg/ha) against Ae. vigilax.

In response to the above, this chapter aims to utilise small-scale field trials to investigate the efficacy of: • VectoBac 12AS and VectoBac WG against Cx. annulirostris in ground-pool freshwater habitats, and • VectoBac G at low rates against Ae. vigilax in saltmarshes.

One of two methods can be used to assess the efficacy of a product in the field. After the water is treated, with a measured amount of insecticide, the mortality of either a) caged (e.g. Brown et al. 1998b) or b) free-swimming mosquito larvae (e.g. Su and Mulla 1999, Amalraj et al. 2000, Fillinger et al. 2003) can be assessed. A pilot trial was conducted to compare the accuracy of these two methods with VectoBac G. Field efficacy of Bti 73

Table 3.1: Label rates of three Bacillus thuringiensis var. israelensis formulations (VectoBac) for application against mosquito larvae in Australia and the USA.

Registered Label Rate Converted to ITU/La

Product Australia USA Australia USA

VectoBac G 7 kg/ha 2.5 – 10 lbs/acre or 933 380 – 1506

2.84 – 11.3 kg/ha

VectoBac 12AS 0.6 – 1.2 L/ha 0.25 – 2 pts/acre or 518 – 1036 240 – 1920

0.3 – 2.4 L/ha

VectoBac WG 125 – 500 g/ha 50 – 400 g/acre or 250 – 1000 246 – 2000

125 – 1000 g/ha

Notes a. Calculated assuming a standard water depth of 15 cm for saltmarsh and mangroves.

3.2 Methods

3.2.1 Development of appropriate sampling techniques

To compare the use of caged or free-swimming larvae, a small-plot trial was conducted. Tests were carried out in round plastic ponds (1.1 m dia., 0.22 m deep) filled with 15 cm of sea water. The sea water (142 L for each pond) was sourced from Moreton Bay, south-east Queensland, Australia. Each pond was lined with 5 cm of washed river sand (Howarths Home and Garden, Duffield Rd, Clontarf). One open-topped, cylindrical bioassay cage (250 µm mesh, 0.5 m dia., 0.2 m deep) was placed in the centre of each pond. The cages were attached to stakes that had been firmly pushed into the sand. The ponds were placed on a flat open area exposed to sunlight at Redcliffe City. The subsequent trials, where the efficacy of Bti was determined in the field, were conducted at Redland Shire and Logan Shire (Figure 3.1).

Caged and free-swimming, laboratory-reared, third-instar Ae. vigilax (colonised from Victoria Point in 2000) were exposed to 4 replicates of 2 rates (533 and 800 Bti ITU/L equivalent to 4 and 6 kg/ha, respectively) of VectoBac G (200 Bti ITU/mg). One hundred larvae were placed into each bioassay cage and 1,000 free-swimming larvae were placed into each pond (Figure 3.2). Larvae were allowed to acclimatise for 1 h before treatment. Ponds were treated with the required amount of product, based on the above rates. Product was applied evenly over the surface of each pond. The product was not applied inside the bioassay cage to ensure that a high concentration of product did not accumulate within the mesh cage, due to reduced water flow.

For each rate, the mortality of the target larvae was recorded after 48 h. Four untreated control ponds were also assessed. To test duration of control, a new cohort of third-instar Ae. vigilax were introduced into the cages and ponds after 5 days (100 added to the cages and 1,000 added to the ponds). As with the initial cohort, mortality was assessed 48 h later (7 days after the treatment was made).

The mortality of caged larvae was calculated after counting the number of surviving larvae inside each cage. The survival of the free-swimming larvae was assessed with a 250-ml capacity dip. One dip was taken at 10 equidistant points around the perimeter of the pond. Field efficacy of Bti 75

Redcliffe

Redland

Logan

300 km Queensland

30 km

Figure 3.1: Map of Queensland and the Moreton Bay region in south-east Queensland (Lat. 27ºS, Long. 153ºE) showing the locations used to examine the efficacy of Bti in the field. 76

Construction 15 cm sea water X 12

5 cm washed river sand

24 h settling period

1000 3rd instar Aedes vigilax 100 3rd instar Aedes vigilax

Treatment 1 h acclimatisation period

Add one treatment from: 4 x VectoBac G per rate 4 x VBC Bti exp. per rate 4 x Untreated control

10 dips taken at separate points

Sample after Number of surviving larvae counted 48 h NOTE: After sampling all exposure larvae were returned.

Figure 3.2: Schematic of experimental design used to compare the sampling techniques of cages and dips used to sample third-instar Aedes vigilax. Field efficacy of Bti 77

The number of larvae in each dip was recorded, and larvae were returned to the pond (i.e. sampling with replacement). A minimum of 10 minutes was allowed between dips for larvae to re-surface.

A Horiba (Kyoto, Japan) portable field laboratory was used to measure the abiotic water characteristics (pH, temperature, salinity). Measurements were taken in all treated and untreated control ponds immediately before treatment, after 48 h, and at the completion of the trial (7 days).

3.2.2 Efficacy of Bti in the field

Based on results from above, larvae were exposed to Bti in mesh cages. Each trial was conducted in replicated ground-pools (freshwater and saltmarsh), containing an open- topped, 250 µm cylindrical mesh cage. One hundred laboratory reared third-instars were placed in each treated and untreated control cage, ca. 2 h beforehand. The number of surviving larvae after 48 h was counted, and the percentage mortality calculated.

A Horiba (Kyoto, Japan) portable field laboratory was used to measure the abiotic water characteristics (pH, temperature, salinity, dissolved oxygen and turbidity [nepholometric turbidity units (NTUs)]). Measurements were taken in all treated and untreated control pools immediately before treatment, after 48 h, and at weekly intervals until completion of the trial.

3.2.2.1 Freshwater ground-pools

The efficacy of VectoBac WG and VectoBac 12AS was assessed in freshwater ground- pools. The trial was conducted from February – June 2002 in Logan Shire, south-east Queensland (Figure 3.1). One cage (0.5 m dia., 0.5 m deep) was placed into the centre of each pool (1 m dia., 0.8 m deep) and adjacent pools were separated by 10 m.

The target species was Cx. annulirostris and 100 laboratory-reared larvae (colonised from Boondall in 1997) were placed in each cage. The caged larvae were exposed to 5 replicates of 6 rates of VectoBac WG: 12, 24, 48, 93, 188 and 390 Bti ITU/L (31, 62, 125, 250, 500 78 and 1,000 g/ha) and 3 rates of VectoBac 12AS: 48, 96, 192 Bti ITU/L (0.3, 0.6 and 1.2 L/ha), in independent pools. Concurrently, 3 untreated control pools were also assessed.

The ponds were treated by adding the appropriate weight of VectoBac WG (weighed in the laboratory with microbalance scales [Mettler Toledo AT261 DeltaRange]) or volume of VectoBac 12AS (measured with a pipette) to 5 L of sieved habitat water. This was then stirred with a glass rod until homogeneous and then poured evenly across the surface of the pool.

To test duration of control, 100 larvae were added to the cages on a weekly basis, until no significant difference in mortality was recorded between untreated and treated pools. As with the initial cohort, mortality was assessed after 48 h.

3.2.2.2 Saltmarsh pools

The efficacy of VectoBac G against Ae. vigilax at various rates was assessed in saltmarsh pools. The trial was conducted from April – May 2003 in Redland Shire, south-east Queensland (Figure 3.1). Caged larvae were exposed to 4 replicates of 133, 266, 533 and 933 Bti ITU/L (1, 2, 4 and 7 kg/ha) of VectoBac G in isolated saltmarsh pools, separated by 30 to 100 m. The concentrations of product were calculated assuming a standard water depth of 15 cm in the saltmarsh. Concurrently 4 untreated control pools were also assessed. Two cages (0.5 m dia., 0.15 m deep) were placed in each replicate pool. The various dosages rates were calculated from the surface area to be treated for each pond; product was weighed in the laboratory with microbalance scales (Mettler Toledo AT261 DeltaRange) and applied by hand. The trials were conducted after the saltmarsh was inundated by the peak of the spring high tides.

3.2.3 Statistical methods

The percent control of caged or free-swimming larvae was calculated, with Abbott’s formula (Abbott 1925):

(c − t ) Percent control = × 100 c Field efficacy of Bti 79 where c = the average number of larvae recorded from untreated control ponds based on n replicates and t = the average number of larvae recorded from treated ponds based on n replicates. Abbott’s formula provides a convenient method for adjusting for natural mortality; however, it does not provide an estimate of variance for the corrected value. Rosenheim and Hoy (1989) present a method for calculating confidence limits for the corrected control value as follows:

(1 − t )× (1 − g ) Proportion control = 1 − ()1 − c

0.5 t ()1 − g  Var()t  ()1 − t 2 × Var()c  Proportion control ± × ()1 − g   +  1 − c  n  2 ()  t  ()1 − c × nc 

Var (c )× e 2 where: g = 2 ()1 − c × nc and e is the critical value chosen from the t distribution, with the desired probability and n – 1 degrees of freedom.

There is no statistical method available to compare different treatments once adjusted with Abbott’s formula. Therefore, the different treatments were compared with a two-way ANOVA using the numbers of deaths recorded per cage (i.e. uncorrected data). To normalise the data, the square root of each proportion was arcsine transformed (Anscombe 1948). Where figures and means are presented, the data have been back-transformed (Anscombe 1948, Zar 1999). Differences in the abiotic water characteristics of the replicate ponds were compared with a one-way ANOVA for each treatment rate. Where necessary, an all-pairwise comparison test (Tukey 1953, cited in Zar 1999) was used to determine significant differences for each factor. 80

3.3 Results

3.3.1 Development of appropriate sampling techniques

The mean total number of third-instar Ae. vigilax recovered from the caged and free- swimming populations is compared in Figure 3.3. The recovery of the caged population was high; between 84 and 90 (out of 100) larvae were recovered from control ponds. In comparison, the recovery of the free-swimming population was much lower and variable; between 15 and 80 (out of 1,000) larvae were recovered from control ponds. The rate of recovery of the free-swimming population was dependant on the weather at the time of sampling. Higher numbers (i.e. 79 – 80) were recovered at times with low wind or rain, when the larvae tended to be clustered around the perimeter of the pools. However, in high wind or rainfall, experienced at day 7, the recovery of larvae was low (i.e. 15 – 31) as larvae were dispersed throughout the water column and the pond.

The mortality of the caged larvae, exposed to both treatment rates, was significantly higher than the untreated controls at 2 days (ANOVA: F1,13 = 199.95, p <0.0001), but not at 7 days (ANOVA: F1,12 = 2.85, p = 0.117).

Due to the change in efficacy of the dip sampling technique, results obtained at one sample period could not be directly compared to another without reference to the number of control larvae. Thus Abbott’s formula was used to compare treatment efficacy (Table 3.2). The caged and dipping methods both indicated that VectoBac G provided effective control at 2 days. However, at 7 days, results were disparate; samples of larvae from the cages indicated no residual control with VectoBac G, yet with the dipping method there appeared to be residual control for the 6 kg/ha treatment (82.95% Abbott’s adjusted control). However, the apparent residual control was not significant due to the low sampling efficacy and the large confidence intervals which eroded the basis for qualitative comparison (the mean [± SD] number of larvae from control ponds was 15 ± 5.89 and treated ponds was 3.25 ± 1.5). It is most likely that the apparent residual control is a false result associated with the difficulties of dip sampling during high winds and rainfall.

The water during the trials was characterised by: warm temperature (range: 28.6 – 32.6ºC), high salinity (range: 2.3 – 3.32%), and neutral pH (range: 6.93 – 8.18). Field efficacy of Bti 81

a) Caged larvae 100

sampled 60

Aedes vigilax 0 Control VectoBac G Control VectoBac G

b) Free-swimming larvae 120 100 80 60 40 20 Mean total number of third-instar 0 Control VectoBac G Control VectoBac G

2 days 7 days

Treatment regime structured by time period for assessment

533 Bti ITU/L (4 kg/ha) 800 Bti ITU/L (6 kg/ha)

Figure 3.3: Mean (± SD) number of Aedes vigilax sampled at 48 h post-treatment from a) cages and b) 10 replicate 250 ml dips of free-swimming larvae. Arcsine data are back transformed.

Table 3.2: Abbott’s adjusted per cent control of third-instar Aedes vigilax exposed to VectoBac G (200 Bti ITU/mg) at various treatment rates for 48 h.

Rate Time post treatment

a 7 days (95 % CL) Bti ITU/L kg/ha 2 days (95 % CL)

Caged larvae 533 4 87.51 (77.01 – 98.02) 3.04 (0.00 – 13.34) 800 6 100.00 (100.00 – 100.00) 14.90 (0.00 – 30.91)

Free-swimming larvae 533 4 100.00 (100.00 – 100.00) 49.26 (25.82 – 72.69) 800 6 100.00 (100.00 – 100.00) 82.95 (71.58 – 94.33)

Notes a. CL = confidence limit.

Field efficacy of Bti 83

3.3.2 Efficacy of Bti in the field

3.3.2.1 Freshwater ground-pools

The trial at Logan Shire was conducted over 5 time periods. As there was no significant difference in the mortality of untreated control larvae from the different times (ANOVA:

F13,28 = 0.67, p = 0.77), it is unlikely that the trials were biased. The water during the trial was characterised by: high turbidity (range: 33 – 525 NTUs), warm temperature (range: 15.8 – 27.1oC), low to medium oxygen content (range: 1.33 – 10.02 g/L), low salinity (range: 0 – 0.01%) and neutral pH (range: 6.40 – 7.77). No statistically significant effect on abiotic water characteristics over time was recorded as a consequence of either VectoBac WG or VectoBac 12AS application.

Mortality of third-instar Cx. annulirostris was compared at 2 sample times: 2 and 9 days post treatment (after each cohort of larvae had been exposed for 48 h). The Abbott’s adjusted mean control of Cx. annulirostris exposed to VectoBac WG and VectoBac 12AS at the various sample times is presented in Table 3.3. Both products provided effective control at 2 days, yet neither provided control at 9 days.

The mortality of Cx. annulirostris after 2 days exposure to VectoBac WG was different between the varying treatment rates, including untreated controls (ANOVA: F6,66 = 34.40, p <0.001). The arcsine back-transformed average (± SD) mortality of untreated control larvae was recorded at 10.43 ± 1.86%, significantly lower than all treatment rates (post hoc: p <0.05). Mortality of larvae exposed to VectoBac WG ranged from 85 ± 3.6% to 100.0 ± 0% in pools treated at rates between 12 and 390 Bti ITU/L (Figure 3.4). At this time, efficacy of the 12 Bti ITU/L concentration was significantly lower than the 48, 188 and 390 Bti ITU/L treatments (post hoc: p <0.05). VectoBac WG treatment efficacy was significantly lower at 9 days (ANOVA: F1,66 = 957.19, p <0.001). At this time, poor control was observed and mortality ranged between 4.0 ± 6.9% and 14.2 ± 12.6%. Untreated control mortality remained low at 12.2 ± 9.7% and was not significantly different from the treated mortality (post hoc: p >0.05).

With respect to VectoBac 12AS, at 2 days there was a statistical difference in the mortality recorded from treated and untreated pools (ANOVA: F3,34 = 89.31, p <0.001). Untreated 84

Table 3.3: Abbott’s adjusted per cent control of caged third-instar Culex annulirostris exposed to VectoBac WG (3,000 Bti ITU/mg) and VectoBac 12AS (1,200 Bti ITU/mg) at various treatment rates after 48 h.

Rate Time post treatment

Bti ITU/L Amount/haa 2 days (95% CL)b 9 days (95% CL)

VectoBac WG

12 31 83.19 (78.13 – 88.26) 4.00 (0.00 – 19.12)

24 62 95.80 (91.53 – 100.00) 0.00 (0.00 – 9.94)

48 125 99.10 (98.00 – 100.00) 8.14 (0.00 – 26.62)

93 250 97.76 (92.59 – 100.00) 0.00 (0.00 – 10.97)

188 500 99.78 (99.19 – 100.00) 3.00 (0.00 – 13.40)

390 1000 100.00 (100.00 – 100.00) 0.00 (0.00 – 16.90

VectoBac 12AS

48 0.3 99.12 (97.44 – 100.00) 0.00 (0.00 – 10.42)

96 0.6 98.90 (97.99 – 99.82) 10.03 (0.00 – 25.16)

192 1.2 100.00 (100.00 – 100.00) 0.00 (0.00 – 11.63)

Notes a. VectoBac WG = g/ha; VectoBac 12AS = L/ha. b. CL = confidence limit. Field efficacy of Bti 85

a) VectoBac WG (3000 Bti ITU/mg) 100 Control 12 (31) 80 24 (62) 48 (125) 60 93 (250)

40 188 (500) 390 (1000)

morality (± SD) 20

0 2 days 9 days Cx. annulirostris b) VectoBac 12AS (1200 Bti ITU/mg)

instar 100 -

Control 80 48 (0.3) 96 (0.6)

Per cent third 60 192 (1.2)

0 2 days 9 days Time post treatment

Figure 3.4: Field evaluation of a) VectoBac WG (3,000 Bti ITU/mg) and b) VectoBac 12AS (1,200 Bti ITU/mg) efficacy against Culex annulirostris after 48 h. Units are presented in Bti ITU/L, brackets = g/ha VectoBac WG and L/ha VectoBac 12AS; Arcsine data are back transformed. 86 control mortality was recorded at 9.0 ± 3.7%, significantly lower than all treatment rates (post hoc: p <0.05). There was no significant difference in the mortality recorded from the various treatment rates (post hoc: p >0.05). The mortality of Cx. annulirostris ranged between 99.0 ± 0.7% and 100.0 ± 0% at 2 days post treatment at rates between 48 and 192 Bti ITU/L (Figure 3.4).

Efficacy of VectoBac 12AS treatment was significantly lower at 9 days (ANOVA: F1,34 = 673.64, p <0.001). At 9 days, poor control was observed with mortality ranging between 6.8 ± 6.5% and 19.6 ± 8.4%. Untreated control mortality was 11.8 ± 8.7%, and was not statistically different from treated pools (post hoc: p >0.05).

3.3.2.2 Saltmarsh pools

The trial was conducted over 2 time periods, and there was no significant difference in the mortality of untreated control larvae between the trials (t test: p = 0.11). The water during the trials was characterised by: high turbidity (range: 9 – 68 NTUs), warm temperature (range: 24.8 – 28.6oC), low to medium oxygen content (range: 0.02 – 10.73 g/L), brackish to saline (range: 0.62 – 4.0%) and neutral pH (range: 7.24 – 8.91). No statistically significant effect on abiotic water characteristics was recorded as a consequence of VectoBac G treatment.

Mortality of third-instar Ae. vigilax was assessed after 2 days (after each cohort of larvae had been exposed for 48 h). The Abbott’s adjusted mean control of Ae. vigilax exposed to VectoBac G is presented in Table 3.4. Effective larval control of >99 % was recorded by dosages of 533 Bti ITU/L (4 kg/ha) and above.

The arcsine back-transformed mortality (± SD) of Ae. vigilax recorded at 2 days was between 56.7 ± 28.9% and 100 ± 0% in pools treated at rates between 133 – 933 Bti ITU/L (Figure 3.5). At this time, the efficacy of the varying rates, including controls, was significantly different (ANOVA: F5,31 26.432, p <0.0001). This difference can be explained by the significantly lower efficacy of the 133 Bti ITU/L rate compared to the 266, 533 and 933 Bti ITU/L rates (post hoc: p <0.05). All of the treatments, excepting 133 Bti ITU/L were significantly different from the untreated controls. Mortality recorded from untreated control pools was 19.53 ± 3.0%. Field efficacy of Bti 87

Table 3.4: Abbott’s adjusted per cent control of caged third-instar Aedes vigilax exposed to VectoBac G (200 Bti ITU/mg) at various treatment rates after 48 h.

Rate

Bti ITU/La kg/ha Adjusted control (95% CL)b

133 1 47.32 (5.25 – 89.40)

266 2 88.36 (63.79 – 100.00)

533 4 99.69 (98.98 – 100.00)

933 7 100.00 (100.00 – 100.00)

Notes a. Concentration of product calculated assuming a standard depth of 15 cm in the saltmarsh. b. CL = Confidence limit.

100

80 60

40 Ae. vigilax 20

0 Percent mortality of caged Control 133 (1) 266 (2) 533 (4) 933 (7) Treatment rate: Bti ITU/L (kg/ha)

Figure 3.5: Field efficacy (mean mortality ± SD) of various VectoBac G (200 Bti ITU/mg) treatments against caged Aedes vigilax in saltmarsh pools after 48 h. Arcsine data are back transformed; concentration of product calculated assuming a standard depth of 15 cm in the saltmarsh. Field efficacy of Bti 89

3.4 Discussion

3.4.1 Development of appropriate sampling techniques

Dip sampling is the most common form of larval sampling world-wide, as it provides for rapid assessment of the larvae (e.g. Su and Mulla 1999, Amalraj et al. 2000, Muller et al. 2001, Fillinger et al. 2003). Miura et al. (1970) provide a preliminary assessment of the technique, who recorded that the technique provides for rapid assessment of population structure (i.e. ratios of instars and pupae) and that sampling time was proportional to the number of larvae collected.

As larvae are not uniformly distributed throughout pools, sampling efforts should be stratified. The small-scale distribution of larvae inside saltmarsh pools was defined by Dale et al. (1986), who recorded that larval distribution was related to water movement. Larvae were recorded to be concentrated on the edges of isolated pools after the tide had receded, particularly along edges with vegetation or towards the drainage outlet of the pond. Field technicians account for such variability by sampling only the edges of ponds (Su and Mulla 1999, Amalraj et al. 2000, Muller et al. 2001), as was undertaken during this trial.

This trial clearly demonstrated that the dip sampling technique is highly variable and dependent on the weather (i.e. wind and rainfall). Varying numbers of larvae sampled from control ponds with dippers have previously been recorded (Miura et al. 1970, Amalraj et al. 2000); however, this is the first trial to directly relate the results to the ambient weather conditions and a known number of larvae inside the ponds.

Due to the changing sensitivity of the dip sampling technique, it was not possible to directly compare the densities of larvae between trials sampled at different times, without standardising the numbers against controls sampled at the same time. This can be achieved with Abbott’s formula (Abbott 1925). Once results are standardised against control values, there remains no statistical method to compare the results and they must be interpreted subjectively. Furthermore, it is not possible to relate the number of larvae per dip to absolute numbers of larvae per water body. The method can only provide a ratio of high or low numbers of larvae for samples taken at the same point in time. However, it does 90 present a useful technique for rapid assessment of the presence or absence of larvae from a water body, particularly when undertaking pre-treatment surveys.

There is a general perception that cages may result in a treatment rate that is higher (if product is trapped inside) or lower (if product is excluded by the mesh) than the surrounding water. This trial indicates that if product is applied outside of the cages, there may be a slightly lower concentration of product inside the cages, as suggested by the lower mortality of caged larvae compared to free-swimming larvae at 7 days. Similarly, Lawler et al. (1999) recorded higher mortality of larvae outside cages than within, after a treatment of VectoBac G at 5.6 kg/ha in the USA. However, this difference is more likely to be related to the low number of free-swimming larvae recovered at this time. When the efficacy of a treatment is assessed with the dip method, it is suggested that results are analysed with caution. It is clearly evident that assessment of the mortality of caged larvae provides a more robust analysis of population mortality. Considering these results, the efficacy of Bti treatments during the field trials was assessed using caged larvae.

3.4.2 Efficacy of Bti in the field

3.4.2.1 Freshwater ground-pools

VectoBac WG was an effective control agent against Cx. annulirostris in freshwater ground-pools. The test concentrations used in the trials were calculated after measuring the depth of the water in the ponds to be 80 cm. This study demonstrates that presenting the results as a concentration opposed to an amount/surface area treated provided the best indication of product use, considering that it is dispersed throughout the water column. In previous small-plot field trials, Ragoonanansingh et al. (1992) and Fillinger et al. (2003) calculated product concentration assuming a standard water depth of 10 cm, but their ponds were actually deeper, thus overestimating product concentration.

Our results indicate that VectoBac WG loses little efficacy when diluted to concentrations equivalent to laboratory LC95 values in the field (see Chapter 2). The registered label rate for VectoBac WG is 125 – 500 g/ha or 48 – 188 Bti ITU/L in 80 cm deep water and for VectoBac 12AS, 0.3 – 1.2 L/ha or 48 – 192 Bti ITU/L in 80 cm deep water. For treatment Field efficacy of Bti 91 of mosquito habitats at depths greater than this, the concentration of product applied at registered label rates, would be approaching those doses where sub-optimal control was observed in the laboratory (Chapter 2; Table 2.1). When selecting an appropriate application rate for mosquito habitat, in particular for freshwater areas, the depth of the water should be considered.

In line with other data, the activity of Bti was lost inside 1 week, and residual activity was not increased with higher concentrations of product (Karch et al. 1991, Kroeger et al. 1995, Fillinger et al. 2003). The present study also confirms that Bti products are benign in terms of abiotic water quality. Based on these results, Bti appears to be an excellent choice for treatment of freshwater ground-pools and loses little efficacy in the field.

Of importance to local operators is the cost comparison of products available for use. A cost ratio of the minimum VectoBac WG and VectoBac 12AS label rates indicates a price ratio of WG:12AS at 1:1, if equivalent volumes of water are used to dilute the product (approximately AUD $27/ha applied with a Bell 47 helicopter). Given that problems with mixing large volumes of VectoBac WG for Ultra Low Volume applications have been indicated previously (Muller et al. 2001), the amount of water may need to be increased for proficient mixing, thus impacting on ferrying time and overall efficacy of aerial treatments. However, VectoBac WG has shown advantages over VectoBac 12AS, not only because of its increased potency against a wide range of mosquito vectors, but also because VectoBac WG can be economically transported, stored and has an extended shelf life.

3.4.2.2 Saltmarsh pools

VectoBac G proved to be effective for the control of Ae. vigilax when applied below the registered label rate in saltmarsh pools. The registered label rate for VectoBac G is 7 kg/ha or 933 Bti ITU/L in 15 cm deep water (Table 3.1). The results herein, indicate that the product is likely to start failing at 266 Bti ITU/L (2 kg/ha) against Ae. vigilax in saltmarsh pools. When the concentration of Bti required for Ae. vigilax control is directly compared with the results for Cx. annulirostris, it is evident that a higher concentration is required in saltmarshes and mangroves. This may be a result of the different environmental characteristics, particularly salinity, the target species, or even formulation characteristics. 92

However, the results in Chapter 2 indicate that both species are highly susceptible to Bti formulations.

There is a paucity of published information on the use of VectoBac G (n = 3). In south-east Queensland, Ritchie (1994) recorded 89.5% control of Ae. vigilax when VectoBac G was applied at 399 Bti ITU/L, and 98.6% control at 933 Bti ITU/L. In the same area, Jeffery et al. (2005) recorded 98% control of Ve. funerea at 933 Bti ITU/L. In Bangladesh, Ali et al. (2000) recorded 76% control of Cx. quinquefasciatus when VectoBac G was applied at 246 Bti ITU/L. These results are similar to those in this study, indicating that formulation characteristics may play a large role in product efficacy.

A limitation of VectoBac G is its low potency, which results in expensive treatments as large amounts of product must be carried to treat a small area. Reducing the application rate from 7 kg/ha to 5 kg/ha would provide a cost saving of almost 30% (price reduced from approximately AUD $63/ha to $45/ha applied with a Bell 47 helicopter). The possibility of reducing the label rate for operational usage is supported by the results of this chapter, as VectoBac G was effective at rates as low as 2 kg/ha. As such, the application rate of 5 kg/ha (or 666 Bti ITU/L in 15 cm deep water) was selected for a large scale field trial to assess product distribution from rotary-wing aircraft (Chapter 4). This rate should provide a considerable safety margin for application error, in terms of control, while also being economical. 93

Chapter 4

Spatial distribution of aerially applied granular Bacillus thuringiensis var. israelensis (VectoBac G)

4.1 Introduction

Over three-quarters of the quantity of insecticide applied for mosquito control is distributed aerially, as the area of saltmarsh and mangrove habitat that requires treatment is extensive and difficult to access by ground. The primary product that has been used for aerial applications during the last 10 years in Australia is liquid Bti (VectoBac 12AS and Teknar). However, granular formulations are preferred over liquids because they can penetrate vegetation canopies more effectively, there is less drift of product, and no loss of chemical due to evaporation (Feng and Sidhu 1989, Ghimire 1997). The use of granular Bti formulations is rapidly gaining popularity and over 12,000 kg of VectoBac G has been applied per year in south-east Queensland since 2003 (CLAG 2005, NEMMO 2005, Figure 1.1). VectoBac G was effective for the control of Ae. vigilax below the registered label rate of 7 kg/ha in Chapter 3. This result implies that the amount of product applied by contractors could be reduced, providing a considerable cost saving to local government control programs.

The effectiveness of an aerial application depends on the correct amount of insecticide being applied, as well as the evenness of the deposition to the target area. It is common to attain the correct average application rate, but have an uneven distribution across the treated area (Ghimire 1997). Areas with under-application may result in sub-optimal control and an increased risk of the larvae developing resistance to the products, while over-application results in wasted product and thus is uneconomical (Bouse and Carlton 1985). Hence, it is 94 important to measure the spatial distribution of aerial applications to ensure that the target area is being correctly treated.

The methodologies used to assess the uniformity or spatial distribution of aerially applied material have been standardised by the American Society of Agricultural Engineers (ASAE Standard 2004). After the applicator flies over systematically arranged catch-trays, the amount of product captured per tray is weighed or counted and the distribution pattern is extrapolated. The coefficient of variation (CV) represents the uniformity of the application and the ASAE standard (2004) suggests that the threshold of acceptability is a CV of less than 20%. Considering the expense and logistics involved with conducting such field trials, it is imperative that trials are well designed and that the size of catch-trays used is appropriate.

In Australia, the methodologies used by contractors to assess the spatial distribution of granular applications have been based on published trials conducted for Altosand, where 1 m2 catch-trays were used (Woods et al. 1996, Woods and Dorr 1997). However, the particle size of VectoBac G (3 – 8 mm dia., 5 – 40 mg) is larger than that of Altosand (0.5 – 1 mm dia., 0.13 – 2.29 mg) and because particle size directly relates to the number of granules available to fall into catch-trays, this has implications for quantitative assessment. Specifically, with a larger particle size, the number of particles available to fall into a catch- tray is less and the variation between replicate catch-trays may be naturally higher (Lockett 1998). To examine the sampling efficiency of different sized catch-trays, the theoretical distribution of particle counts expected per catch-tray was modelled, based on the Poisson distribution. Using this information, the optimum size of catch-trays was calculated for VectoBac G and this was compared with Altosid pellets, which have a larger particle size (3 – 120 mm dia., 10.9 – 560 mg) and Altosand, which has a smaller particle size.

The results of the modelling procedure provided a basis for designing a field trial to evaluate if the application rate of VectoBac G could be lowered to 5 kg/ha (or 666 Bti ITU/L in 15 cm deep water) and using aerial applicators control Ae. vigilax and achieve a uniform distribution of product. This constitutes the first quantitative assessment of an aerial VectoBac G application in Australia. During the experimental treatment, the mortality of caged larvae placed in isolated pools was assessed, in addition to measuring the deposition of VectoBac G in catch-trays. Aerially applied Bti 95

The common method used to visualise the spatial distribution of product is to plot deposition against the geographical position of the catch-tray and interpolate between the sampling points. To achieve such sophisticated analysis, spatial programs such as ArcView GIS, can now be used to produce contour plots. Contour maps have been used to visualise the spatial distribution of granular temephos and s-methoprene formulations as well as liquid Bti formulations (Woods et al. 1996, Woods and Dorr 1997). The utility of spatial programs as a support tool for analysing product distribution was also examined. 96

4.2 Methods

4.2.1 Modelling product distribution

A simulated stochastic-modelling procedure was used to calculate the distribution of product that would be theoretically expected after an aerial treatment, for granules of different sizes (VectoBac G, Altosid pellets and Altosand). This information was used to determine the optimum size of catch-trays for evaluating aerially applied granular products. Firstly, assuming that the product is distributed evenly to the site, the theoretical expected number of particles per catch-tray was calculated with the formula:

v × y z PART = x where z PART = number of particles; v = application rate (g/m2); y = catch-tray size (m2) and x = mean particle mass (g) based on mass data from 200 individual particles.

Next, a distribution pattern was constructed around z PART. The distribution pattern was based on the Poisson distribution; however, if particle counts were large, i.e. greater than 100, the normal distribution was employed. The Poisson distribution is useful for describing random occurrences, when the probability of occurrence is small; however, as the probability of occurrence increases, the Poisson distribution becomes symmetrical to the normal distribution (Zar 1999). To create a distribution pattern that could be statistically compared to observed data, a cumulative frequency distribution was built. The number of particles expected per catch-tray 95% of the time was calculated by removing the tails of the curve (probability less than 0.025 and above 0.975).

Considering that technical operations are conducted with the units of kg/ha, z PART was converted to a weight per catch-tray. This was achieved by simulating the number of particles per catch-tray 1,000 times (based on the above distribution pattern). Each simulated particle was randomly assigned a weight from 200 particles weighed in the laboratory using microbalance scales (Mettler Toledo AT261 DeltaRange). Using these modelled data, the theoretical distribution pattern of mass expected in catch-trays of specific sizes was built. Aerially applied Bti 97

The theoretical weight of product per catch-tray was simulated 1,000 times for a range of catch-tray sizes at set application rates (5 kg/ha for VectoBac G, 4 kg/ha for Altosid pellets and 4 kg/ha for Altosand). The simulated data were used to calculate the upper and lower 2.5% bounds of probability, indicating the bounds of which 95% of simulated points fall within. When graphed, the width of these points indicated the amount of sampling error associated with each specific catch-tray size.

The optimal sample size (number of catch-trays) was investigated by modelling the expected mass per tray 100 times for different sample sizes (2 to >200 as required). Using these data, the 95% confidence interval that surrounded the estimate of the mean was calculated, compared graphically with sample size, and the number of catch trays required to estimate the mean application rate with ± 5% accuracy was identified.

4.2.2 Aerial trials

The aerial trial evaluated the spatial distribution of VectoBac G, applied at 5 kg/ha, as well as the efficacy of the treatment to control Ae. vigilax. The trial was conducted in a saltmarsh at Redland Shire Council (Figure 3.1), and the product was distributed with a rotary-wing aircraft (Bell 47, McDermott Aviation Pty. Ltd.) fitted with twin hoppers and calibrated to disperse VectoBac G at the nominated rate (Figure 4.1). The contractor had carried out broad-scale applications of pesticides on a regular basis for over 10 years and selected the nominated flight lane separation (14 m).

The product dispensed during the trial was sampled in 43 replicate 1 m2 catch-trays arranged in a grid-like pattern in the target area. The catch-tray size of 1 m2 was selected as it was the nominated size used by contractors for quality assurance of aerial granular applications (Woods et al. 1996, Woods and Dorr 1997). The catch-trays were placed near the mangrove and saltmarsh boundary, with half of the catch-trays under mangrove canopy. The position of the catch-trays and flight path were recorded with a dGPS (Trimble System). The product deposited into each catch-tray was removed and transported to the laboratory, where it was weighed using microbalance scales (Mettler Toledo AT261 DeltaRange) and the number of particles was counted. 98

Figure 4.1: VectoBac G (200 Bti ITU/mg) was distributed with rotary-wing aircraft, fitted with specially designed hoppers for product dispersal (Photo Credit: Robert A. Fusco, Valent Biosciences). Aerially applied Bti 99

The efficacy of the treatment was assessed against laboratory-reared, third-instar Ae. vigilax. The larvae were placed in open topped, 250 µm cylindrical mesh cages (0.50 m dia. by 0.20 m deep), held in place by stakes and circular mesh support, ca. 2 h beforehand. Eight cages, each containing 100 larvae, were placed in isolated saltmarsh pools separated by 10 to 90 m, and at least 5 cm in depth, in both the treatment site and an adjacent untreated control site. The number of surviving larvae was counted at 48 h post-treatment.

4.2.3 Statistical methods

The amount of VectoBac G deposited in catch-trays in the open and under canopy was compared with a t test (“Student” 1908). The Kolmogorov-Smirnov test (Kolmogorov 1933, Smirnov 1939a, b, cited in Zar 1999) was used to determine if the probability distribution of the observed number of particles per tray differed from the theoretical cumulative frequency distribution.

The dGPS positions of the catch-trays and flight path were downloaded and overlaid on an aerial photograph of the site using ArcView GIS 3.2a (Environmental Systems Research Institute [ESRI] 1996). The standard method to graphically analyse the distribution pattern is to interpolate between the sample points to produce a contour map (Woods et al. 1996, Woods and Dorr 1997). The conversion of point observations into a surface is based upon a user-defined interpolation technique and kriging has been identified in the literature as the best linear unbiased estimator (Isaaks and Srivastava 1989). The use of an interpolation technique, such as kriging, is based on the assumption that the observed weights at a fixed number of locations can be mathematically modelled.

Moran’s I analyses were used to assess the degree of spatial autocorrelation, or interdependence, in the weight of VectoBac G collected in catch-trays separated by different distances (Cliff and Ord 1981). Moran’s I analyses have been widely used for the study of spatial patterns (Basso et al. 2001, Ryan et al. 2004, Jung et al. 2006), and in this study Moran’s I was used to determine whether weights of VectoBac G were randomly distributed, or whether there was a significant clustering of catch-trays with high or low weights. Analyses were based on 3 lag classes (i.e., catch-tray separation distances with mutually exclusive intervals): 0 – ≤10 m, >10 – ≤20 m and >20 – ≤30 m. The boundaries of 100 these intervals were chosen so that each lag class had at least 30 pairs of catch-trays, and the maximum lag class was less than one-half the minimum dimension of the study area. The weights of VectoBac G were transformed to the natural log to homogenise the variance. Values closer to 1.0 indicate that catch-trays within that lag class were positively correlated; values closer to –1.0 indicate that catch-trays within that lag class were negatively correlated; and values around 0 indicate that there is no autocorrelation and the spatial distribution of weights was random. The spatial autocorrelation of correlograms was tested using a Monte Carlo simulation method and adjusted for repeated sampling with a Bonferroni correction (Howell 1997). Spatial autocorrelation analyses were conducted using Rookcase software (Sawada 1999).

Universal kriging methods were used to estimate the weight of VectoBac G at unsampled locations throughout the study area. Kriging is a method of spatial prediction that incorporates a model of the covariance of the random function and uses a weighted moving average interpolation to produce the optimal spatial linear prediction (Cressie 1991). VectoBac G weights were interpolated at 1 x 1 m intervals. A linear drift function was used to model the underlying trend. Local averaging was based on a fixed search radius of 60 m around each interpolated point, unless there were <8 catch-trays within 60 m. In this case, the radius was expanded to include a minimum of 8 catch-trays. Analyses were conducted using ArcView GIS 3.2a (ESRI 1996).

To determine whether the universal kriging method provided reliable estimates of VectoBac G weights, a leave-one-out cross-validation analysis was used. This involved the stepwise removal of individual catch-trays and the remaining points were used to predict the weight that would be collected. This process was repeated for each catch-tray and the expected and actual data were compared using a linear regression. A strong linear association between observed and predicted weights, with a slope close to 1.0, would indicate that the weight of VectoBac G in catch-trays was adequate to explain the variability between catch-trays. Conversely, a weak linear association between actual and predicted weights of VectoBac G would indicate that catch-trays near each other were not closely related. In this case, the data were not interpolated and presented as a contour map; rather these data were presented as a bubble map, without interpolation (Fleischer et al. 1999). Aerially applied Bti 101

The distance from each catch-tray to the nearest flight path was measured in ArcView GIS 3.2a (ESRI 1996). A Spearman Rank correlation (Spearman 1904) was used to analyse the relationship between the weights of VectoBac G captured in each tray and the distance to the nearest flight path.

The mortality of larvae exposed to the treatment was statistically compared with the untreated control larvae using a t-test (“Student” 1908) on the numbers of deaths recorded per cage. To normalise the data, the square root of each proportion was arcsine transformed (Anscombe 1948). Where means are presented, the data have been back-transformed (Anscombe 1948, Zar 1999).

102

4.3 Results

4.3.1 Modelling product distribution

Based on the Poisson distribution, the expected number of particles per catch-tray was calculated for VectoBac G, Altosid pellets and Altosand at different application rates and catch-tray sizes (Table 4.1). Using this information, cumulative frequency distributions were built to provide a graphical means of comparing theoretical data with experimental data (Figure 4.2). For VectoBac G and Altosid pellets – which have a relatively large granule size – the amount of product in each catch-tray was relatively low. With pellets in particular, there was a 10% chance that no product would be captured in a 1 m2 catch-tray at the registered label rate (3 – 4 kg/ha). Furthermore, it was not possible to clearly discriminate between the 95% boundaries of probability of the expected number of particles for different treatment rates. Using VectoBac G as an example, when applied at 5 kg/ha, the theoretical expected number of particles (z PART) was 25, and 95% of counts were expected to be between 15 and 36 particles. At 7 kg/ha, z PART was 35 and 95% of counts were between 24 and 48 particles. The effect of increasing the catch-tray size when assessing VectoBac G and Altosid pellets was to increase the number of particles expected per tray, but it also increased the ability to discriminate between the bounds of different treatment rates. For example, when assessing VectoBac G applied at 5 kg/ha with 2 m2 catch-trays, z PART was 50, and 95% of counts would be between 37 and 65 particles. At 7 kg/ha, z PART was 71 and 95% of counts would be between 55 and 88 particles.

To provide modelling information that was in the technical units of kg/ha, the theoretical number of particles expected per catch-tray was converted to a weight per catch-tray. The proportional relationship between catch-tray size and sampling accuracy was also evident when the 95% bounds of the weight per catch-tray was converted to kg/ha and compared (Table 4.2). This was supported graphically: a) in a series of graphs that can be directly compared with experimental data in technical units (Figures 4.3 to 4.5) and b) when the upper and lower 2.5% boundaries of probability were modelled for Altosid pellets and Altosand at 4 kg/ha and for VectoBac G at 5 kg/ha (Figure 4.6). Altosid pellets had the lowest sampling efficacy; for example when targeting 4 kg/ha, the mass of product captured with a 1 m2 catch-tray would be expected to vary by the equivalent of 10.6 kg/ha. Increasing the catch-tray size to 4 m2 reduced the variation in expected weight to 4.93 kg/ha. VectoBac Aerially applied Bti 103

Table 4.1: The 95% bounds calculated for the expected number of particles per catch-tray of different sizes for Altosid pellets (40 g AI/kg), VectoBac G (200 Bti ITU/mg) and Altosand (4 g AI/kg).

Treatment rate Catch-tray size (m2) (kg/ha) 1 2 3 4 Altosid pellets 3 0 – 6 1 – 9 2 – 12 4 – 15 4 0 – 6 2 – 11 4 – 15 6 – 20

VectoBac G 5 15 – 36 37 – 65 60 – 94 83 – 122 7 24 – 48 55 – 88 88 – 127 119 – 167

Altosand 3 269 – 338 556 – 651 844 – 960 1143 – 1267 4 364 – 438 746 – 858 1137 – 1274 1530 – 1686 104

2 a) Altosid pellets in 1 m d) Altosid pellets in 4 m2 1 1

0.8 0.8

0.6 0.6

0.4 0.4

0.2 0.2

0 0 0 5 10 15 20 0 10203040

2 b) VectoBac G in 1 m e) VectoBac G in 2 m2 1 1

0.8 0.8

0.6 0.6

0.4 0.4

0.2 0.2 C u m 0 0 u l a 0 20 40 60 80 100 0 20 40 60 80 100 120 140 t i v e Number of particles per catch tray p 2 r o c) Altosand in 1 m b a 1 b i l i t y 0.8 2 kg/ha 3 kg/ha 0.6 4 kg/ha 0.4 5 kg/ha 6 kg/ha 0.2 7 kg/ha 8 kg/ha 0 0 100 200 300 400 500 600 700 Number of particles per catch tray

Figure 4.2: The cumulative frequency Poisson distribution of the expected number of particles of Altosid pellets (40g AI/kg; 2 – 5 kg/ha), VectoBac G (200 Bti ITU/mg; 4 – 8 kg/ha) and Altosand (4g AI/kg; 2 – 6 kg/ha) in different sized catch-trays at different treatment rates. Note: scales on x axis are different. Aerially applied Bti 105

Table 4.2: The width of the theoretical 95% bounds for the treatment rate (kg/ha) calculated from the simulated weight per catch-tray, in trays of different sizes for Altosid pellets (40 g AI/kg), VectoBac G (200 Bti ITU/mg) and Altosand (4 g AI/kg).

Treatment rate Catch-tray size (m2) (kg/ha) 1 2 3 4 Altosid pellets 3 8.47a 6.30 4.88 4.20 4 10.60 7.64 5.82 4.93

VectoBac G 5 4.83 2.79 2.46 2.00 7 4.60 3.48 2.61 2.35

Altosand 3 0.75 0.55 0.44 0.38 4 0.87 0.61 0.52 0.46

Notes a. Upper 95% bound minus lower 95% bound (kg/ha). 106

a) 4 kg/ha in 1 m2 f) 4 kg/ha in 2 m2 20 20

15 15

10 10

5 5

0 0

b) 5 kg/ha in 1 m2 g) 5 kg/ha in 2 m2 20 20

15 15

10 10

5 5

0 0

2 c) 6 kg/ha in 1 m2 h) 6 kg/ha in 2 m 20 20

15 15

10 10

5 5

0 0

Percentage of catch-trays d) 7 kg/ha in 1 m2 i) 7 kg/ha in 2 m2 20 20

15 15

10 10

5 5

0 0

2 e) 8 kg/ha in 1 m 2 j) 8 kg/ha in 2 m 20 20

15 15

10 10

5 5

0 0 0.01 - 0.05 0.16 - 0.20 0.31 - 0.35 0.46 - 0.50 0.61 - 0.65 0.76 - 0.80 0.91 - 0.95 1.06 - 1.10 1.21 - 1.25 1.36 - 1.40 1.51 - 1.55 1.66 - 1.70 1.81 - 1.85 1.96 - 2.00 2.11 - 2.15 0.01 - 0.05 0.16 - 0.20 0.31 - 0.35 0.46 - 0.50 0.61 - 0.65 0.76 - 0.80 0.91 - 0.95 1.06 - 1.10 1.21 - 1.25 1.36 - 1.40 1.51 - 1.55 1.66 - 1.70 1.81 - 1.85 1.96 - 2.00 2.11 - 2.15

Mass of VectoBac G (g) per catch-tray

Figure 4.3: The theoretical distribution of the mass of VectoBac G (200 Bti ITU/mg) in 1 and 2 m2 catch-trays at different treatment rates (4, 5, 6, 7 and 8 kg/ha). Aerially applied Bti 107

a) 2 kg/ha in 1 m2 e) 2 kg/ha in 4 m2 25 25 20 20 15 15 10 10 5 5

0 0

2 b) 3 kg/ha in 1 m f) 3 kg/ha in 4 m2 25 25

20 20

15 15

10 10

5 5

0 0

c) 4 kg/ha in 1 m2 g) 4 kg/ha in 4 m2 25 25

20 20

15 15 Percentage of catch-trays 10 10

5 5

0 0

2 d) 5 kg/ha in 1 m2 h) 5 kg/ha in 4 m 25 25 20 20 15 15 10 10 5 5 0 0 0.01 - 0.10 0.31 - 0.40 0.61 - 0.70 0.91 - 1.00 1.21 - 1.30 1.51 - 1.60 1.81 - 1.90 2.11 - 2.20 2.41 - 2.50 2.71 - 2.80 3.01 - 3.10 3.31 - 3.40 3.61 - 3.70 3.91 - 4.00 0.01 - 0.10 0.31 - 0.40 0.61 - 0.70 0.91 - 1.00 1.21 - 1.30 1.51 - 1.60 1.81 - 1.90 2.11 - 2.20 2.41 - 2.50 2.71 - 2.80 3.01 - 3.10 3.31 - 3.40 3.61 - 3.70 3.91 - 4.00

Mass of Altosid pellets (g) per catch-tray

Figure 4.4: The theoretical distribution of the mass of Altosid pellets (40 g AI/kg) in 1 and 4 m2 catch-trays at different treatment rates (2, 3, 4 and 5 kg/ha). 108

a) 2 kg/ha 50 40

30 20 10

b) 3 kg/ha 50 0.976 - 1.00 0.01 - 0.025 0.901 - 0.925 0.826 - 0.850 0.751 - 0.775 0.675 - 0.700 0.601 - 0.625 0.526 - 0.550 0.451 - 0.475 0.376 - 0.400 0.301 - 0.325 0.226 - 0.250 0.151 - 0.175 40 0.076 - 0.100 30 20 10

c) 4 kg/ha 50 0.976 - 1.00 0.01 - 0.025 0.901 - 0.925 0.826 - 0.850 0.751 - 0.775 0.675 - 0.700 0.601 - 0.625 0.526 - 0.550 0.451 - 0.475 0.376 - 0.400 0.301 - 0.325 0.226 - 0.250 0.151 - 0.175 0.076 - 0.100 40

30 20

0 Percentage of catch-trays d) 5 kg/ha 50 0.976 - 1.00 0.01 - 0.025 0.901 - 0.925 0.826 - 0.850 0.751 - 0.775 0.675 - 0.700 0.601 - 0.625 0.526 - 0.550 0.451 - 0.475 0.376 - 0.400 0.301 - 0.325 0.226 - 0.250 0.151 - 0.175 40 0.076 - 0.100 30

20 10 0

e) 6 kg/ha 50 0.976 - 1.00 0.01 - 0.025 0.901 - 0.925 0.826 - 0.850 0.751 - 0.775 0.675 - 0.700 0.601 - 0.625 0.526 - 0.550 0.451 - 0.475 0.376 - 0.400 0.301 - 0.325 0.226 - 0.250 0.151 - 0.175 40 0.076 - 0.100 30 20

10 0 0.976 - 1.00 0.01 - 0.025 0.901 - 0.925 0.826 - 0.850 0.751 - 0.775 0.675 - 0.700 0.601 - 0.625 0.526 - 0.550 0.451 - 0.475 0.376 - 0.400 0.301 - 0.325 0.226 - 0.250 0.151 - 0.175 0.076 - 0.100

Mass of Altosand (g) per catch-tray

Figure 4.5: The theoretical distribution of the mass of Altosand (4 g AI/kg) in 1 m2 catch-trays at different treatment rates (2, 3, 4, 5 and 6 kg/ha). Aerially applied Bti 109

a) Altosid pellets at 4 kg/ha 12

0 12345678 Catch tray size (m2) b) VectoBac G at 5 kg/ha 10 9 8 7 6 5 4 3 2 1 0 12345678

Bounds for theoretical application rate Catch tray size (m2) c) Altosand at 4 kg/ha 6

2 pp

0 0.25 0.5 0.75 1 1.25 1.5 1.75 2 2.25 2.5 2.75 3 Catch tray size (m2)

Boundaries95% confidence of probability limit 50% Inter-quartile confidence limit range

Figure 4.6: The theoretical bounds of product distribution for a) Altosid pellets (40 g AI/kg) at 4 kg/ha, b) VectoBac G (200 Bti ITU/mg) at 5 kg/ha and c) Altosand (4 g AI/kg) at 4 kg/ha, calculated from the simulated mass captured in catch-trays of different sizes. Note: scales on x axis are different. 110

G also had a low sampling efficacy when using 1 m2 catch-trays, as the mass of product captured would be expected to vary by 4.83 kg/ha when targeting 5 kg/ha. Increasing the catch-tray size to 2 m2 reduced the variation in the expected weight to 2.79 kg/ha. In practical terms, the recommended catch-tray size was defined as the point in Figure 4.6 where increasing the catch-tray size further did not significantly reduce the theoretical variation in product catch, when the increased workload and infrastructure is considered. The recommended catch-tray size for VectoBac G was 2 m2 and for Altosid pellets was 4 m2.

In contrast, Altosand, which has a smaller granule size, can be sampled with a 1 m2 catch- tray with high efficacy. High numbers of particles are expected to be recovered in a 1 m2 catch-tray (95% bound = 364 – 438 particles for 4 kg/ha) and there is a clear distinction in the 95% bounds of different treatment rates (Table 4.1, Figure 4.2 and 4.5). With a tray size of 1 m2, the simulated mass of product would be expected to vary by the equivalent of only 0.87 hg/ka when targeting 4 kg/ha.

As the number of catch-trays used to estimate the mean application rate is increased, the 95% confidence intervals for the calculated mean decrease (Figure 4.7, Table 4.3). Also, more catch-trays were required to sample products that had larger granules with the same level of accuracy (error = ± 5% of application rate). For example, the number of catch-trays required to estimate the mean with ± 5% accuracy for Altosid pellets (4 kg/ha in 4 m2) was 159, VectoBac G (5 kg/ha in 2 m2) was 35 and Altosand (4 kg/ha in 1 m2) was 4.

Aerially applied Bti 111

7.0 a) Altosid pellets at 4 kg/ha

6.0

5.0

4.0

3.0

2.0

1.0 0 5 10 15 20 25 30 35 40 45 50 55 60 65 70

7.0 b) VectoBac G at 5 kg/ha

6.5

6.0

5.5

5.0

4.5

4.0

3.5 9

5 3.0 % 0 5 10 15 20 25 30 35 40 45 50 55 60 65 70 C o n f i

d 4.3 c) Altosand at 4 kg/ha e n c

e 4.2 i n t e 4.1 r v a l s 4.0 ( k g

/ 3.9 h a ) 3.8

3.7

3.6 0 5 10 15 20 25 30 35 40 45 50 55 60 65 70 Sample size (n )

Catch-tray size: 1 m2 2 m2 3 m2 4 m2

Figure 4.7: The 95% confidence intervals surrounding the mean application rate (kg/ha) calculated from 100 simulations for each sample size (n) for a) Altosid pellets (40 g AI/kg) at 4 kg/ha, b) VectoBac G (200 Bti ITU/mg) at 5 kg/ha and c) Altosand (4 g AI/kg) at 4 kg/ha. 112

Table 4.3: The number of catch-trays required to evaluate an aerial application of Altosid pellets (40 g AI/kg), VectoBac G (200 Bti ITU/mg) and Altosand (4 g AI/kg) in catch-trays of different sizes to achieve an estimate of the mean with ± 5% accuracy.

Treatment rate Catch-tray size (m2) (kg/ha) 1 2 3 4 Altosid pellets 3 765 416 292 212 4 698 360 216 159

VectoBac G 5 70 35 33 16 7 45 27 12 12

Altosand 3 2 2 2 2 4 4 2 2 2 Aerially applied Bti 113

4.3.2 Aerial trials

During the trial, 15 ha of saltmarsh were treated at 10.00 h on the 27th November 2003. At the time of treatment there was a southerly wind blowing at 3.5 m/s, the ambient air temperature was 23.5ºC and the relative humidity was 60%. The aircraft flew north to south transects, parallel to the wind, in a progressive (back-and-forth) flight path.

The mean (± SD) treatment rate of VectoBac G calculated from the mass of product deposited in 1 m2 catch-trays was 5.76 ± 3.46 kg/ha (CV = 60%), just above the target rate. The average treatment rate was slightly higher in the open area at 6.08 ± 3.36 kg/ha (CV = 60%) and lower under the canopy at 5.26 ± 3.21 kg/ha (CV = 70%); however, there was no statistical difference between the deposition in open and canopy areas (student’s t test: p = 0.45). The coefficient of variation of 60 – 70% indicated that the product was not distributed evenly to the target area.

The results from the aerial trial were overlaid on the theoretical distribution patterns (section 4.3.1) to compare the experimental data with the simulated model data (Figures 4.8 and 4.9) and it is evident that the product was not distributed evenly over the site. The number of VectoBac G particles captured in 1 m2 catch-trays was significantly greater than the Poisson distribution for 5 kg/ha (Kolmogorov-Smirnov test: T = 0.746, p <0.01).

The mean (± SD) flight lane separation was 14.70 ± 4.52 m (calculated by measuring the distance between consecutive flight paths at 10 m intervals in ArcView GIS 3.2a, ESRI (1996). This was close to the targeted lane separation of 14 m; however, the variation in lane separation was large. As such, only 25.83% of the area was covered with a lane separation between 12 and 16 m. When the flight path was graphically examined (Figure 4.10), these changes in lane separation were clearly evident.

The standard method for visualising the spatial distribution of product is to interpolate between the sample points to produce a contour map. Prior to this, Moran’s I analyses of the weight of VectoBac G captured in catch-trays was used to determine whether there was spatial autocorrelation in catch-tray weights. A weak positive autocorrelation of catch-trays within 10 m was evident, however, at greater distances there was no spatial correlation between catch-trays (Table 4.4). The accuracy of the universal kriging method to predict 114

1 y

t 0.8 i l i b a b

o 0.6 r p e v i t 0.4 a l u m u 0.2 C

0 0 10 20 30 40 50 60 70 80 90 Number of VectoBac G particles per 1 m2 catch tray

5 kg/ha Poisson distribution 7 kg/ha Poisson distribution

Actual data

Figure 4.8: The cumulative frequency distribution of VectoBac G (200 Bti ITU/mg) particles captured in 1 m2 catch-trays, in the saltmarsh, after the aerial treatment in March 2003 (n = 26), compared with the Poisson distribution expected when treating at 5 and 7 kg/ha (see Figure 4.2).

Aerially applied Bti 115

20 a) 5 kg/ha Observed 15 Theoretical

0 Percentage of catch trays

20 b) 7 kg/ha Observed 2.51 - 2.55 2.41 - 2.45 2.31 - 2.35 2.21 - 2.25 2.11 - 2.15 2.01 - 2.05 1.91 - 1.95 1.81 - 1.85 1.71 - 1.75 1.61 - 1.65 1.51 - 1.55 1.41 - 1.45 1.31 - 1.35 1.21 - 1.25 1.11 - 1.15 1.01 - 1.05 0.91 - 0.95 0.81 - 0.85 0.71 - 0.75 0.61 - 0.65 0.51 - 0.55 0.41 - 0.45 0.31 - 0.35 0.21 - 0.25 0.11 - 0.15 0.01 - 0.05 15 Mass of VectoBac G per 2 m2 catch tray Theoretical Percentage of catch-trays 10

0 Percentage of catch trays 2.51 - 2.55 2.41 - 2.45 2.31 - 2.35 2.21 - 2.25 2.11 - 2.15 2.01 - 2.05 1.91 - 1.95 1.81 - 1.85 1.71 - 1.75 1.61 - 1.65 1.51 - 1.55 1.41 - 1.45 1.31 - 1.35 1.21 - 1.25 1.11 - 1.15 1.01 - 1.05 0.91 - 0.95 0.81 - 0.85 0.71 - 0.75 0.61 - 0.65 0.51 - 0.55 0.41 - 0.45 0.31 - 0.35 0.21 - 0.25 0.11 - 0.15 0.01 - 0.05 Mass of VectoBac G per 1 m2 catch tray

Figure 4.9: The distribution of VectoBac G (200 Bti ITU/mg) mass captured in 1 m2 catch- trays, in the saltmarsh, during the aerial treatment in March 2003 (n = 26), compared with the theoretical distribution of expected mass at a) 5 kg/ha and b) 7 kg/ha (see Figure 4.3). 116

50 metres

b) 1 2 3 4 5

Weight (g) of VectoBac G per catch tray (weight classes are based on theoretical 95% bounds of mass per tray) (Underdosed) (Target rate) (Overdosed) (Numbered at top)

50 metres

Figure 4.10: Position of bioassay cages, catch-trays and helicopter flight path that a) is overlaid on an aerial photograph of Redlands Shire Council, and b) displays the weight of VectoBac G (200 Bti ITU/mg) captured in each catch-tray after the treatment in March 2003 (Photo credit: Redland Shire Council). Aerially applied Bti 117

Table 4.4: Moran’s I spatial autocorrelation indices for weights of VectoBac G collected in catch-trays after the aerial treatment in March 2003.

Lag distance (m) Moran’s I p-value 0 – ≤10 m 0.2142 0.027 * >10 – ≤20 m – 0.0075 0.394 >20 – ≤30 m – 0.1035 0.104

Notes The spatial autocorrelation of correlograms was tested using a Monte Carlo simulation method and adjusted for repeated sampling with a Bonferroni correction. * p <0.05 118 weight of VectoBac G at unsampled locations (i.e. by using the leave-one-out cross- validation method) was low. Regression analysis of the relationship between predicted versus observed weights was not significant (R2 = 0.086, p = 0.071). This indicates that contour mapping, using kriging methods, was not a reliable method for analysing VectoBac G distribution.

The amount of product deposited in the grid of catch-trays was spatially displayed with a bubble plot (Figure 4.10). The theoretical 95% bounds of mass per tray (0.300 – 0.725 g; section 4.3.1) were used when developing the bubble plot. There was no clear pattern in the spatial distribution of weight and only 54% (n = 24) of the catch-trays were treated at the target rate (within the theoretical 95% bounds; 18% [n = 8] underdosed and 28% [n = 11] overdosed). The presence of canopy did not seem to affect the amount of product caught, and 53% (n = 9) of catch-trays under canopy were treated at the target rate (23.5% [n = 4] underdosed and 23.5% [n = 4] overdosed), which is comparable to catch-trays in the open that were treated at the target rate 58% (n = 15) of the time (15% [n = 4] underdosed and 27% [n = 7] overdosed). There was one row of catch-trays (directly east of flight-path 4 in Figure 4.10) where 7 out of the 8 catch-trays were over-treated; however, there does not appear to be any obvious relationship with the position of the flight-path (Figure 4.10) or the presence of canopy (50% were under canopy) when this row in particular is examined.

When the weight of product captured in each catch-tray is compared with the distance to the nearest flight path, there is a parabolic (non-linear) relationship, as shown in Figure 4.11. Catch-trays close to the flight-path (<2 m) caught relatively low amounts of product, which can be related to the action of the spreader (Figure 4.1), which distributes granules with a sideward velocity. Relatively high amounts of product were caught in catch-trays that were between 2 and 6 m from the helicopter, and at distances over 6 m the weight of product captured was relatively low. Due to this pattern, a Spearman Rank correlation (cf. linear regression) was used to analyse the relationship between distance to the nearest flight path and the weight captured in each tray, and there was no significant relationship (p >0.05).

The mean (± SD) mortality of caged Ae. vigilax at 48 h post-treatment was 100.0 ± 0.0% in pools subject to the aerial treatment of VectoBac G. Eight control cages were originally constructed for analysis; however, during the 48 h period proceeding the trial, high wind- gusts were experienced and consequently five control cages were blown over, leaving only Aerially applied Bti 119 three for analysis. At 48 h post-treatment, mortality of untreated control larvae was recorded at 26.0 ± 0.1%. The mortality of the larvae in the treatment area was statistically higher than that recorded in the control pools (Student’s t-test: p <0.0001). The catch-trays were positioned according to the availability of isolated pools and were close to trays that caught the target amount of product (Figure 4.10). 120

1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 Weight of VectoBac G (g) 0 0246810 Distance to nearest flight path (m)

Figure 4.11: Weight (g) of VectoBac G (200 Bti ITU/mg) captured per catch-tray compared with the distance to the nearest flight path (m) after the aerial treatment in March 2003 targeting 5 kg/ha.

Aerially applied Bti 121

4.4 Discussion

Previously, aerial contractors had conducted quality assurance for distribution of all granular products based on methodologies published for Altosand, where 1 m2 catch-trays were used (Woods et al. 1996, Woods and Dorr 1997). Although 1 m2 catch-trays can accurately sample Altosand, based on theoretical analysis, they cannot accurately sample VectoBac G and Altosid pellets. This was directly related to the larger particle size of VectoBac G and Altosid pellets. In the USA, Bouse and Carlton (1985) also found 1 m2 catch-trays to be too small for analysis of the distribution pattern of large herbicide pellets (1 cm3), and catch-trays that were 3.7 by 6 m were used for greater accuracy. If trials are conducted to assess VectoBac G and Altosid pellets with 1 m2 catch-trays, the wide bounds of the expected number and weight of product per tray must be taken into consideration when analysing the trial, as was done when constructing the bubble plot (Figure 4.10). Based on the results of the modelling procedure, it is recommended that tray size be at least 4 m2 for Altosid pellets, 2 m2 for VectoBac G and 1 m2 for Altosand. Little gain in precision would be made by increasing the catch-tray size further, when the increased workload and infrastructure is considered.

The optimum number of catch-trays that would be required for accurate sampling was also influenced by the particle size of the products. Due to the high accuracy that can be achieved when sampling Altosand, a low number of catch-trays (2 – 4) would be required to estimate the mean application rate with 95% accuracy. However, to provide useful results on the spatial distribution of any product, Nansen et al. (2003, 2006) has suggested that at least 50 catch-trays is required. This should be taken into account when selecting the number of catch trays from Table 4.3.

The results from the modelling procedure (section 4.3.1) were used to produce graphic and tabular reference material that could be used by council operators and contractors when conducting quality assurance trials. When designing field trials, the first step would be to select the optimum size of catch-trays. Then, the optimum number of catch-trays can be selected from Table 4.3 (keeping in mind the minimum requirement of 50 trays for spatial analysis). After the experimental data have been collected, they can be graphically compared with the theoretical expected distribution in Figures 4.2 to 4.6. This is a quick and 122 simple procedure that provides useful information on how uniform the distribution of product was and where the mean application rate lies.

Further information about the spatial distribution can be provided by producing a surface map. However, experimental data can have many distributions (i.e. normal, skewed or random) and this will affect whether kriging can accurately interpolate between sample points to produce a contour map (Isaaks and Srivastava 1989). Thus, it is important to conduct an a priori analysis, to test for spatial continuity between samples, over distances that exceed the distance between each sample, before producing a spatial map (Nansen et al. 2006). The results of this trial indicate that the widely accepted contour mapping (Wood et al. 1996, Woods and Dorr 1997) was not appropriate for VectoBac G and that bubble mapping provided more useful information.

Even though the modelling procedure indicated that 1 m2 trays were not optimal for assessing VectoBac G, it was necessary to demonstrate the inaccuracy inherent in the current operational monitoring system before improvements could be made. During the field trial, the distribution of product to the site was uneven, and this may have resulted from a number of factors, including uneven flight lane separation, accuracy of the dGPS system, wind gusts, accuracy of sampling technique (catch-trays as discussed above), and variability in particle size.

Although wind gusts occurred, wind direction was made more or less in line with the designated flight path so this may have had minor effect. Wind gusts have the potential to skew the distribution of product to one side of the helicopter, creating patches of over and under application (Sutherland et al. 1974). It is difficult for the helicopter pilot to adjust the flight path to account for the effects of wind gusts. Therefore, it is usual for applications to be made at times of low wind speed. The wind speed experienced during the trial (3.5 m/s) was low and unlikely to have affected product distribution.

Variability in particle size has the potential to introduce additional error with regard to the uniformity of application (Yates et al. 1973, Bansal et al. 1998, Fulton et al. 2005). This was first noted by Yates et al. (1973) who used equations of motion to predict the lateral spread of particles released from aircraft. The terminal velocity of a particle is related to the size of the particle, which in turn affects lateral spread. Consequently, products such as VectoBac Aerially applied Bti 123

G and Altosid pellets, which have a large and variable particle size may be more subject to vagaries in distribution within a swath width.

While the spatial distribution of product was easily defined, it is more difficult to evaluate the biological consequences of variability in product deposition. Despite the uneven application in the trial, there was 100% control of the caged Ae. vigilax. However, 18% of the area was underdosed, receiving the equivalent of between 0.5 to 2.84 kg/ha. According to the results in Chapter 3, it is likely that control of field larvae would have ranged between 47.32 to 88.36% in the area that was underdosed; however, bioassay cages were not placed in these areas. Even though the application of VectoBac G at 5 kg/ha was uneven, it still provided a sufficient safety margin for satisfactory treatment of Ae. vigilax.

Being able to apply VectoBac G at 5 kg/ha would provide operators with a cost saving of almost 30%, when compared with the registered label rate of 7 kg/ha. From this trial it is evident that a uniform distribution of VectoBac G is difficult to achieve, even under benign weather conditions. The most important factors that affected the observed spatial distribution of product were the uneven flight path of the helicopter and the low sensitivity of the small catch-trays.

When resources allow, a trial using 2 m2 catch-trays should be initiated, and the results communicated to the contractor in order to remove what seemed to be a major confounding variable in quantitative assessment of application efficacy. It is recommended that the swath pattern of VectoBac G is characterised from single passes of the helicopter over a line of 2 m2 catch-trays. This information could be used to calculate the optimal lane separation of the helicopter for multiple-pass applications and used as base-line information for designing a multiple-pass trial. It is likely that routine broad-scale applications of VectoBac G are no better (or worse) than what was achieved in this trial. Apart from the potential for sublethal dosing, with implications for resistance, over-treatment may bring with it additional risk of environmental impact. These issues have been explored in Chapter 5.

125

Chapter 5

Evaluation of potential ecological impacts of Bacillus thuringiensis var. israelensis and s-methoprene on non- target organisms of saltmarsh and mangrove habitats

5.1 Introduction

Despite the fact that Bti and s-methoprene are considered to be among the most target specific of insecticides (e.g. Ali 1981, Mulla et al. 1982, Brown et al. 2000a; see section 1.7), there are indications that non-target organisms may be impacted in different ecosystems (e.g. lentic freshwater: Norland and Mulla 1975, Hershey et al. 1998; lotic freshwater: Car and de Moor 1984, Back et al. 1985). Many laboratory studies have provided information on the specific responses of different taxa to Bti and/or s-methoprene, with the majority of these studies recording no impacts when non-target organisms were exposed to operational concentrations (see section 1.7). However, laboratory-based trials fail to account for ecological processes, such as individual stress and population-level interactions, which may alter the effects of the pollutant in the field (Ward et al. 1976, Underwood 1995, Chapman 2002). Only one study, by Hershey et al. (1998), has investigated the long-term ecological impacts of Bti and s-methoprene under field conditions. The results of this work contradicted those from many laboratory-based toxicological experiments showing that the density of freshwater insects was significantly reduced for 3 years during repeated applications of Bti and s-methoprene. Clearly, there is a need for more field-based ecological testing on the impacts of these insecticides.

Furthermore, although there is a basic understanding of the impacts of Bti and s-methoprene on non-target organisms in freshwater systems in the northern hemisphere, it may be inappropriate for managers to apply this information to Australian saltmarsh systems (Adam 126

2002). The tropical and subtropical saltmarshes along the east coast of Australia present a unique ecosystem that are colonised by a high diversity of relatively short grasses and are found in close proximity to mangroves, with the mangroves forming a barrier between the saltmarsh and the sea (Adam 2002). In these regions, the saltmarshes are only inundated by the highest spring high tides (Adam 1994, Morrisey 1995). Australian saltmarshes provide many ecosystem services, including a high level of primary production and inputs into coastal food webs (Coull et al. 1995, Bouillon et al. 2002, Mazumder et al. 2006), habitat for fishes (Morton et al. 1987, 1988, Connolly 1999, Thomas and Connolly 2001) and shorebirds (Loyn et al. 1986, Major 1991), and a buffer zone between the land and the sea (Bridgewater and Cresswell 1999, Faulkner 2004). There is, however, almost no information on the ecological impacts of Bti and s-methoprene used for mosquito control in Australian saltmarshes, despite this being the primary means by which the transmission of pathogens from mosquitoes to humans is controlled (Russell and Kay 2004).

In this chapter, the potential for applications of Bti and s-methoprene to affect the density and diversity of non-target invertebrates in subtropical Australian saltmarshes was investigated. During mosquito control, the insecticides are applied as a blanket to the saltmarsh; thus, both terrestrial and aquatic organisms are exposed. Therefore, the trial was designed to sample both the terrestrial and aquatic invertebrates inhabiting the saltmarsh. The results of this paper provide essential information for mosquito control operators and environmental managers, to minimise the human health problems associated with saltmarshes in the most environmentally sound manner possible.

Ecological impact assessment 127

5.2 Methods

5.2.1 Study area

The study was carried out in three locations (Pine Rivers, Coomera Waters and Garden Island), in Moreton Bay, south-east Queensland, Australia (Figure 5.1), which is a large subtropical, estuarine embayment. The intertidal flats at the localities are characterised by 3 broad zones: a saltmarsh fringe along the upper shore (primarily Sarcocornia quinqueflora and Sporobolus virginicus and inter-dispersed with saltpans), a mid-intertidal mangrove flat (primarily Avicennia marina) and a lower-intertidal mud flat with intermittent sea grass beds (primarily Zostera capricorni) (Adam 1994). All locations were typical of sites that would be targeted for mosquito control in south-east Queensland (Dale et al. 1986) and were at least half a hectare in size and without significant grazing by hoofed animals. The locations had previously been treated for mosquito control, primarily with Bti, at a similar intensity each summer, ca. once per month at the recommended label rate. At the time of the experiments the locations had been free from insecticide use for at least 1 month (local council records).

Tides at the localities are semidiurnal with a range of 1.2 to 2.7 m (Pine Rivers) and 0.8 to 2.0 m (Coomera Waters and Garden Island) within a full monthly tidal cycle. The mudflat and mangroves are inundated on a daily basis by high tides, whereas the saltmarsh is inundated for up to four consecutive days during spring tides (flooded for an average of 4 – 5 h per high tide) but rarely on other occasions. On a yearly basis, about 7% of tides totally inundate the saltmarsh, although this varies across seasons, with more inundations during the day in the Austral summer and during night in the Austral winter (Connolly 2005). As the water drains from the saltmarsh with the receding tide, water remains in depressions in the substratum, forming stagnant pools that persist for 1 – 7 days.

During the study, the effect of experimental insecticide applications on non-target terrestrial and aquatic arthropods was examined in the saltmarsh. Terrestrial arthropods are the most dominant form of saltmarsh fauna and include insects, mites and spiders (Moreton et al. 1988, Morrisey 1995; Table 5.1). The aquatic invertebrates that inhabit saltmarshes include nematodes, molluscs (bivalves and gastropods) and crustaceans (crabs, prawns, shrimp,

128

Pine Rivers

Garden Island

Coomera 300 km Queensland Waters W t

30 km

Figure 5.1: Map of Queensland and the Moreton Bay region in south-east Queensland (Lat. 27ºS, Long. 153ºE) showing the locations used to examine the effects of Bti and s- methoprene on non-target organisms. Ecological impact assessment 129 amphipods and copepods) (Morgan and Hailstone 1986, Chapman et al. 1998, Breitfuss 2003, Guest and Connolly 2004).

5.2.2 Experimental design

The general hypothesis examined in this study was that applications of Bti and/or s- methoprene to experimental plots would cause the assemblages of non-target arthropods to change in a way that was different from the untreated control plots. Some groups of arthropods (e.g. chironomids) may be adversely affected by the application of insecticides, showing direct, acute impacts (e.g. Hershey et al. 1998). On the other hand, some groups may show indirect effects, including possible increases or decreases in abundance mediated by a change in the abundance of prey, competitors or predators. No indirect effects of Bti and s-methoprene on non-target arthropods have been previously demonstrated, but this is not surprising considering the paucity of field trials that have been conducted.

To test this hypothesis, experimental applications of both Bti and s-methoprene were added to plots in the saltmarsh habitat and over the proceeding 20 days non-target arthropods were sampled from both insecticide-treated and untreated control plots and abundances were compared. The experimental plots (detailed below) were established along a visually uniform stretch of saltmarsh without natural barriers (creeks etc.). The plots were selected to represent the different habitats that are treated with insecticide during routine control operations, when the insecticides are applied as a blanket to the saltmarsh and, as such, plots were positioned in both terrestrial and aquatic habitats.

5.2.3 Development of appropriate sampling techniques

Initially, a pilot experiment was done to develop appropriate methods for sampling terrestrial and aquatic invertebrates in the saltmarsh (Table 5.2). This was necessary as there was little available information on the composition and abundance of terrestrial and aquatic invertebrates in Moreton Bay saltmarshes. It was important to test the validity of the methods under a range of different conditions, due to the ephemeral nature of the pools and the potential for different numbers and types of non-target arthropods to be present at different times of the year and in different places (Morrisey 1995). The pilot experiment 130

Table 5.1: The non-target organisms that have been recorded in subtropical saltmarshes in Moreton Bay, south-east Queensland. Taxa Reference Phylum MOLLUSCA Class Gastropoda Ophicardelus quoyi Morgan and Hailstone 1986, Ophicardelus ornata Breitfuss 2003 Salinator solida Onchidina australis Guest and Connolly 2004 Phylum ARTHROPODA Class Insecta Order Diptera (Nisia nervosa) Morton et al. 1988 Order Lepidoptera (Acrodipsas illidgei) Breitfuss and Dale 2004 Order Coleoptera (Family Hydrophilidae) Morton et al. 1988 Order Hemiptera Morton et al. 1988 Order Hymenoptera (S.F. Formicoidea) Morton et al. 1988 Class Arachnida (Dolomedes sp.) Morton et al. 1988 Class Crustacea Order Amphipoda Morton et al. 1988 Order Decapoda Heloecius cordiformis Chapman et al. 1998, Breitfuss Helograpsus haswellianus 2003, Guest et al. 2006 Parasesarma erythodactyla Fenneropenaeus merguiensis Connolly 2005 Australoplax tridentata Guest et al. 2006 Leander tenuicornis Brown et al. 1996 Subclass Copepoda Canuellidae Coull et al. 1995 Stenhelia sp. Ectinosoma sp. Halicyclops sp. Order Cladocera Morton et al. 1988

Ecological impact assessment 131

Table 5.1 cont. Taxa Reference Phylum CHORDATA Class Osteichthyes Atherinosoma microstoma Connolly et al. 1997, Thomas and Gobiopterus semivestitus Connolly 2001 Torquigener pleurostictus Morton et al. 1987, Thomas and Acanthopagrus australis Connolly 2001 Ambassis marianus Liza argentea Torquigener hamiltoni Valamugil georgii Mugilogobius stigmaticus Connolly 2005 Calamiana sp. Gerres subfasciatus Pseudomugil signifer Brown et al. 1998a, Morton et al. 1998 Class Mammalia Xeromys myoides Van Dyke 1996, Dickman et al. 2000 132

ment). SD) b,c ± n Bay to examine the impacts T:44.0±29.5;T:86.8±54.3; C:34.4±23.9 T: 142.9 ± C:143.3±159.1 132.2; C: 87.1 ± 104.2 T: 144.9 ± 149.0; C:T:42.0±16.9; 123.1 ± 109.8 T: 163.7 ± C:93.4±76.4 144.5; C: 104.3T:72.3±57.2; ± 106.5 C:63.0±67.5 ; mean 2 b b b b a a a 4.0 ± 0.0 Size of plots (m Terrestrial Aquatic of 8 plots per experiment). e, control vegetation and control bare plots; i.e. total of 16 plots per experi plots in the saltmarsh. methoprene that were conducted in three locations in Moreto - s 10Mar2005 GardenIsland 33.0±0.0 16Dec2004 GardenIsland 33.0±0.0 13Jan2005 CoomeraWaters 33.0±0.0 9Feb2004 CoomeraWaters9Feb2005 4.0±0.0 CoomeraWaters 33.0±0.0 Treatment date Location 8Jan2004 PineRivers 9 Mar 2004 Coomera Waters 4.0 ± 0.0 and Bti = 4 for each experimental factor (treated and control plots; i.e. total = 4 for each experimental factor (treated vegetated, treated bar VectoBac 12AS, 1.2 L/ha VectoBac 12AS, 1.2 L/ha n n -methoprene, Altosid Liquid Larvicide, 0.36 L/ha Main experiment Bti, Active Ingredient, Formulation and Treatment rate Pilot experiment Bti, s c. T = Treated, C = Control. b. a. Table 5.2: The experimental treatments of on non-target arthropods sampled from terrestrial and aquatic Ecological impact assessment 133 comprised three experimental applications of Bti in two different locations (Pine Rivers and Coomera Waters) between January and March 2004 (Table 5.2). Two consecutive applications, one month apart, of Bti were done at Coomera Waters. The Pine Rivers locality was only available for one experimental application. Due to its close proximity to urban development, the local government was required to implement broad-scale mosquito control operations after the January 2004 experimental application and this made the location unsuitable for further use during this study.

For each of the three experimental applications, plots were selected within the saltmarsh at each location: 16 terrestrial plots (8 with vegetation [S. quinqueflora and/or S. virginicus] and 8 bare of vegetation [saltpans]; each 2 x 2 m in size) and 8 ephemeral pools (5 to 15 cm deep; Table 5.2). Bti (VectoBac 12AS, Valent Biosciences, 1.2 L/ha) was applied to half of the plots (i.e. 4 vegetation terrestrial, 4 bare terrestrial and 4 ephemeral pools) and the rest were left as untreated controls. The insecticide-treated and untreated plots were randomly inter-dispersed throughout the location. At Coomera Waters, the plots that were selected for the two consecutive insecticide applications were separated from each other by a distance of 500 m, so any effects from the first application would not have affected the plots used for the second.

The terrestrial plots (vegetation and bare) were sampled for non-target invertebrates at 1, 4, 10 and 20 days post-insecticide-application with pitfall traps and aspirators. Pitfall traps have been found to be effective at sampling crabs in saltmarshes (Chapman et al. 1998) and insects and arachnids in grasslands (Standen 2000, Borges and Brown 2003). Aspirators (suction samplers) have been shown to be effective at sampling insects and arachnids from low-standing vegetation (Borges and Brown 2003, Gratton and Denno 2005). Two pitfall traps (33.18 cm2 aperture, 80 mm deep), separated by at least 1 m, were set in each plot, positioned so that the top of the trap was flush with the ground. In the vegetated plots, the pitfall traps were positioned within the strand of S. quinqueflora and/or S. virginicus. In the bare plots, the traps were positioned greater than 1 m from the nearest vegetation. The traps were opened for 24 h before being sampled, with a small amount (10 – 20 ml) of detergent and water (ratio 1:1) placed in the bottom. Two aspirator samples were taken from the vegetated plots. A metal frame (0.5 x 0.5 m) was thrown haphazardly into the plot, to select the position for sampling, and arthropods trapped in the frame were sampled from the vegetation using a hand-held vacuum cleaner (Black and Decker, 3.6V). The collection 134 nozzle sampled an area of 0.001 m2 and was moved to evenly cover the area within the metal frame in 1 minute. Arthropods were contained in a cotton bag placed inside the aspirator.

The ephemeral pools were sampled for non-target invertebrates at 1 and 4 days posttreatment with a sweep net, after this time there was insufficient water remaining in the pools for further samples to be collected. A metal frame (0.5 x 0.5 m) was thrown into the pool, within 10 cm of the edge, pressed into the sediment, and any trapped invertebrates were sampled from the water using a sweep net (10 x 20 cm aperture) with 250 µm mesh, for 1 minute. Two samples were collected from each pool, separated by the maximum distance possible.

The samples were washed across a 250 µm sieve to remove sediment and debris and the contents were then preserved in 70% ethanol. All animals from the terrestrial samples were removed, counted and identified to the lowest taxonomic level possible. Due to large numbers of animals in the aquatic samples, a 10% subsample was sorted. The entire sample was placed in 500 ml of water and stirred with a magnetic stirrer to create a homogenous solution, then 50 ml was extracted, from which all animals (except mosquito larvae) were removed, counted and identified to the lowest taxonomic level possible. In many habitats, it has been found that identification of organisms to the level of species is unnecessary for the detection of environmental impacts (Herman and Heip 1988, Moore and Bett 1989). Therefore, individual taxa were analysed at the taxonomic level of order or subclass. Analysis of these data (see Results) indicated there was a large amount of variation between replicate samples that was likely to be caused by sampling biases, which resulted from the area sampled with the sweep net, methods for employing pitfall traps and the number of replicates needed, for example. The sampling methods were therefore modified to reduce these artefacts in the main experiments (described below).

Differences in the abundance of individual taxa were analysed using a two-way univariate ANOVA with factors for Treatment and Time. Data were transformed to the natural log to meet the assumptions of homoscedasticity of variances after Cochran’s test (Underwood 1981). In all cases, transformation removed heteroscedasticity of variances. When appropriate, post-hoc pooling of mean square estimates (Underwood 1981) was used to determine significant differences for each factor. Ecological impact assessment 135

5.2.4 Selection of experimental plots

An experimental application of each product was done at two different locations to determine whether any impacts were consistent across the region. The Coomera Waters locality, on the western side of the bay, was affected by urban, rural and industrial developments, whereas Garden Island, situated in the centre of the bay, is an undeveloped low-lying island. Thus, the quality of the environment varied considerably between the localities. There was no specific interest in comparing developed to undeveloped areas, only to examine the generality of any effects on non-target saltmarsh fauna.

An experimental application for each of Bti (VectoBac 12AS, Valent Biosciences) and s- methoprene (Altosid Liquid Larvicide [ALL], Sumitomo Chemicals Co.; Table 5.2) was done at each location. The separate applications of each of the insecticides were done at least one month apart and it is unlikely that the results of the successive application were confounded by the previous application because: a) the plots that were selected for each separate insecticide application (i.e. experiment) were separated by a distance of at least 500 m and b) the products have a short activity in the field (Bti is active for 2 – 3 days and s- methoprene is active for 7 days).

For each of the trials, plots were selected within the saltmarsh at each location: 8 terrestrial plots (covered in strands of S. quinqueflora and/or S. virginicus; 11 x 3 m) and 8 ephemeral pools (5 to 15 cm deep) (a total of 64 plots). There were few areas within the terrestrial habitat that were devoid of vegetation and the results of the pilot experiment indicated that there were no significant differences in the abundance of individuals between bare and vegetated plots (see Results), so bare plots were excluded from these trials. The plots were permanently marked with 40 cm long wooden stakes, pushed into the substratum at the corners of terrestrial plots and at one end of each pool. Adjacent plots were separated by 80 to 120 m.

In each trial, four terrestrial plots and four aquatic plots were designated at random for insecticide treatment. The product, diluted with tap water (left to sit for at least 24 h beforehand to allow chlorine to evaporate) at a ratio of 1:2, was applied evenly to the surface of the treated plots at the recommended label rate, using a pressure-pump hand-held sprayer (Hills Garden Sprayer). The remaining four terrestrial and four aquatic plots served 136 as untreated controls. The experimental treatments were done between December 2004 and March 2005 (Table 5.2) between 10.00 and 11.00 h. The insecticide was applied immediately after the spring high tides period when the saltmarshes are flooded, so the experimental plots were not tidally-flooded during the course of the experiment (20 days). The application rate of the chemicals and timing of the trials within the monthly tidal cycle mimicked the mosquito control operations that are regularly carried out by local governments in south-east Queensland.

5.2.5 Sampling technique

The sampling design was developed based on the results of the pilot experiments. To minimise biases leading to increased spatial variability in the estimates of abundance and composition of the dominant fauna (see Results) the area sampled with each technique (pitfall traps, aspirator samples and aquatic sweeps) was increased relative to the pilot experiment.

The terrestrial plots were sampled with pitfall traps and aspirators for non-target invertebrates at 1, 5, 8 and 20 days after the plots were treated. To reduce biases from repeated sampling in the same spot (Underwood 1989, Skilleter 1996), a different 2 x 2 m sub-plot was sampled at each occasion. Four pitfall traps (each consisting of a set of 3 close- set traps with a total aperture of 99.54 cm2, each 80 cm deep) were dug into the ground, flush with the substrate, in the corner of each subplot (30 cm inside). Any gaps around the edges of the trap were packed with mud. The traps were opened for 24 h before being sampled, with a small amount (10 – 20 ml) of detergent and water (ration 1:1) placed in the bottom. After each sampling occasion, the traps were removed from the substratum and the holes filled. One sample of arthropods from the vegetation was taken from each sub-plot using the aspirator (Black and Decker, 3.6 V). The entire 2 x 2 m sub-plot was sampled with the metal frame used in the pilot trial, and one person gradually worked through each sub- plot, walking from one side to the other, moving the aspirator to evenly cover the area of the sub-plot within 1 minute.

The ephemeral pools were sampled for non-target organisms using a sweep net (20 x 30 cm aperture, 250 µm mesh) at 1 and 4 days post-insecticide-application, after this time there Ecological impact assessment 137 was insufficient water remaining in the pools for further sampling. The samples were collected without the metal frame used in the pilot trial, and sweeps were made from the centre of the pool towards the outside, in an area of approximately 1 m2, for 1 minute. Two samples, separated by the maximum distance possible, were collected from each pool.

Samples were processed as described for the pilot experiment, but the aquatic samples were first fixed in 7% formalin containing the stain Rose Bengal, a vital (cell) stain that helps differentiate living from dead animals, and only animals living at the time of collection were counted.

5.2.6 Statistical methods

Initially, the effect of Bti and s-methoprene application on the composition and/or relative abundances of different taxa was compared using multivariate analyses to examine the simultaneous responses of the different taxa that composed the communities. Data on the composition and abundance of non-target invertebrates collected after each experimental insecticide application (total 4 treatments, Table 5.2) and each sampling technique (n = 3) were analysed separately for each occasion (total of 12 analyses). This was necessary as each experimental insecticide application and sampling technique was independent. During each individual analysis, the numbers of non-target arthropods sampled from insecticide- treated and untreated control plots were compared over time. With time the water evaporated from the ephemeral pools and insufficient water remained for further samples to be collected, this occurred at different rates in the pools and in some replicates occurred before samples could be collected on day 4, leaving an unbalanced design. When necessary, replicate pools were randomly selected for inclusion in the statistical analysis to create a balanced design.

Differences in the community composition between insecticide-treated and untreated plots were examined using PERMANOVA, non-parametric multivariate analysis of variance (Anderson 2001), with terms Treatment (fixed: treated versus control), Time (fixed: terrestrial = 4 levels; aquatic = 2 levels), Plot (nested within Treatment x Time, 4 levels) and replicate samples (nested within Plot (Treatment x Time)). The differences were examined graphically using non-metric multidimensional scaling (nMDS) in PRIMER (Clarke 1993). 138

The nMDS ordination ranks the similarity of each pair of samples based on a user-defined similarity measure and visualises the similarity of the communities in three-dimensional space. The stress value associated with each graph was calculated when plotting the three- dimensional graphs on a flat surface. The PERMANOVA and nMDS analyses were based on the Bray-Curtis similarity measure and fourth-root transformed data. The Bray-Curtis similarity measure summarises the species present and the relative abundances of species into a single index that can be used to quantify how assemblages differ through space and time. The fourth-root transformation was used because samples were dominated by a few very abundant taxa (e.g. copepods) and this transformation places greater emphasis on the rare species (Clarke 1993).

The individual taxa that contributed most to the separation of the assemblages in the different treatments were identified with SIMPER analysis. Differences in the abundance of individual taxa that contributed greater than 10% to the community composition were analysed using a two-way univariate ANOVA with factors for Treatment and Time. Data were transformed to the natural log to meet the assumptions of homoscedasticity of variances after Cochran’s test (Underwood 1981). In all cases, transformation removed heteroscedasticity of variances. When appropriate, post-hoc pooling of mean square estimates (Underwood 1981) was used to determine significant differences for each factor. Ecological impact assessment 139

5.3 Results

5.3.1 Development of appropriate sampling techniques

During the pilot experiment, there was considerable variation in the abundance of non-target organisms (copepods) collected from ephemeral pools. Specifically, some samples had extremely large numbers of non-target individuals per 0.25 m2 (e.g. >3000 copepods), while other samples contained no individuals. It is most likely that the metal frame (0.25 m2) used to delineate the pond area for sampling was too small, and swarms of copepods that commonly form during daylight (Ambler et al. 1991, Leising and Yen 1997) were missed. To overcome this, the sampling approach was changed to sample a larger area of the pond without the metal-frame, in an effort to increase the change of sampling swarms.

There was also considerable variation in the abundance of non-target organisms (adult dipterans and hymenoptera) collected in pit-fall traps. Again, some samples contained large numbers of individuals (e.g. >550) while others contained none. Previously, Abensperg- Traun and Steven (1995) demonstrated that increasing the diameter of pitfall traps will increase the abundance of hymenopterans. Thus, to increase the chance of sampling non- target organisms, both the aperture (99.54 cm2) and number (n = 4 per 2 m2) of pitfall traps was increased.

There was low numbers of non-target organisms (adult dipterans and heteropterans) removed from S. quinqueflora and S. virginicus by the aspirator. Specifically, 87.5% of samples contained less than 30 individuals. It was noted that throwing the metal-frame into the vegetation caused a lot of disturbance and many flying insects did not resettle; however, the method did present advantages as it was simple and quick to use in the field. To increase the number of individuals sampled, a larger area was sampled (2 m2) without the metal- frame. 140

Table 5.3: Mean (± SD) total numbers of individuals collected from terrestrial and aquatic plots, after treated plots were exposed to Bti. The percent reduction of individuals in treated plots is highlighted, but there was no statistical difference between treated and control means (ANOVA: p >0.05).

Sample Location Control Treatment technique (treatment date) Day ( x ± SD) ( x ± SD) % Changea Pitfall Pine Rivers 1 33.5 ± 58.0 17.6 ± 25.2 – 48 (08/01/04) 4 12.5 ± 9.4 16.1 ± 22.6 + 28 10 8.3 ± 8.3 13.75 ± 25.6 + 65 Coomera Waters 1 29.8 ± 70.5 16.1 ± 25.9 – 45 (09/02/04) 4 5.3 ± 3.7 15.3 ± 29.2 + 188 10 16.7 ± 32.9 5.4 ± 3.8 – 68 20 17.2 ± 42.3 2.6 ± 2.2 – 85 Coomera Waters 1 10.8 ± 13.2 9.8 ± 9.6 – 9 (09/03/04) 4 18.5 ± 39.7 6.8 ± 3.1 – 64 10 7.6 ± 9.5 10.1 ± 17.0 + 33 20 14.5 ± 41.3 4.4 ± 4.2 – 69

Aspirator Pine Rivers 1 41.25 ± 28.7 27.3 ± 22.7 – 35 (08/01/04) 4 23.6 ± 9.2 22.4 ± 9.5 – 5 10 30.3 ± 14.3 24.3 ± 15.0 – 20 Coomera Waters 1 3.0 ± 1.1 3.4 ± 0.7 – 12 (09/02/04) 4 24.1 ± 27.9 11.3 ± 6.1 – 53 10 15.0 ± 7.6 10.3 ± 10.0 – 32 20 9.9 ± 7.4 7.6 ± 5.4 – 23

Coomera Waters 1 6.4 ± 5.8 3.5 ± 1.4 – 45

(09/03/04) 4 10.0 ± 3.5 13.4 ± 5.7 + 34

10 14.5 ± 6.4 11.9 ± 4.4 – 18

20 19.3 ± 10.0 11.3 ± 6.6 – 42

Aquatic Pine Rivers 1 774.5 ± 810.4 497 ± 585.9 – 35 (08/01/04) 4 525.9 ± 554.8 482 ± 733.4 – 8 Coomera Waters 1 1371 ± 1707.4 1320.4 ± 1510 – 3 (09/02/04) 4 1940 ± 2395.5 1070.3 ± 1632.9 – 44 Coomera Waters 1 2699.5 ± 2673.3 1427 ± 2002.5 – 47 (09/03/04) a. % change = reduction or increase in total number of individuals in treatment plots relative to the control plots sampled at the same time. Ecological impact assessment 141

5.3.2 Impacts of Bti and s-methoprene

5.3.2.1 Effects on community composition

The effects of the experimental treatments on the composition of terrestrial and aquatic taxa were inconsistent between the two localities in south-east Queensland. Specifically, after the Bti treatments, the composition of terrestrial and aquatic communities was different between treated and control plots at Coomera Waters (PERMANOVA, p <0.05, Table 5.4) but only the composition of the aquatic community was different at Garden Island. The effect of Bti on the terrestrial community at Coomera Waters was significant at 5 days post-treatment (pitfalls, post hoc: p = 0.047) and by 8 days the communities from the treated and control plots were similar (post hoc: p >0.05). The timing of the change in the aquatic community varied between the localities and was significant on day 1 at Coomera Waters (post hoc: p = 0.003) and on day 4 at Garden Island (post hoc: p = 0.004).

The different responses of the communities inhabiting the different localities were also detected after the s-methoprene applications. After the s-methoprene applications, the relative composition of terrestrial and aquatic communities between treated and control plots was different at Coomera Waters (PERMANOVA, p <0.05, Table 5.4) but not at Garden Island. The effect of s-methoprene did not interact with time (Table 5.4).

The community responses to Bti and s-methoprene exposure were confirmed graphically with multidimensional scaling (nMDS) and the graphs had small stress values (<0.20). Generally the nMDS graphs did not show a clear separation of the communities on treated and control plots, even when the PERMANOVA recorded a significant result, indicating weak responses of the communities. Due to the large number of plots (n = 40), plots are only presented for communities that had a significant Treatment x Time interaction, after post-hoc pooling of the mean square estimates to identify the times at which the communities were significantly different (Figure 5.2). There was poor separation of the terrestrial community at Coomera Waters after the Bti treatment on day 5. There was a weak separation of the aquatic communities evident after the Bti treatments at both Coomera Waters (day 1) and Garden Island (day 4), and this may be a consequence of the low number of control pools because the water evaporated before samples could be collected. 142

Table 5.4: Differences in community composition between treated (Bti and s-methoprene) and control plots, compared with non-parametric multivariate analysis of variance (using PERMANOVA).

p value

Sample type Treatment Time Interaction

Bti Coomera Waters Aspirator 0.003 * 0.001 * 0.963 Pitfall 0.031 * 0.001 * 0.018 * Sweep Net 0.001 * 0.001 * 0.001 *

Garden Island Aspirator 0.963 0.001 * 0.096 Pitfall 0.695 0.001 * 0.973 Sweep Net 0.025 * 0.001 * 0.006 *

s-methoprene Coomera Waters Aspirator 0.244 0.001 * 0.373 Pitfall 0.007 * 0.001 * 0.059 Sweep Net 0.004 * 0.014 * 0.129

Garden Island Aspirator 0.729 0.063 0.541 Pitfall 0.766 0.001 * 0.031 * Sweep Net 0.768 0.001 * 0.005 *

Notes Data were transformed to the fourth-root and analysis was based on the Bray-Curtis similarity measure. * p <0.05.

Ecological impact assessment 143

Stress:Stress: 0.14 0.14

b) Stress:Stress: 0.01 0.01

c) Stress:Stress: 0.01 0.01

Control

Treatment

Figure 5.2: nMDS ordinations on fourth-root transformed species abundance data based on the Bray-Curtis similarity measure from replicate samples from Bti-treated and control plots. Significant interactions between Treatment x Time were recorded with non- parametric multivariate analysis of variance (Table 5.4) at a) Coomera Waters from terrestrial plots sampled with pitfall traps at 5 days (n = 4 replicates from four treated and control plots), b) Coomera Waters from aquatic pools at 1 day (n = 2 replicates from four treated and three control pools) and c) Garden Island from aquatic plots at 4 days (n = 2 replicates from four treated and two control pools). 144

5.3.2.2 Effects on abundance of taxa

A total of 15 arthropod taxa (2 Arachnida Classes, 4 Crustacea Classes and 9 Insecta Classes) and 1,869,734 individuals were collected and identified during the experiment. After the experimental Bti treatments, two taxa, Acariformes and Copepoda, showed different abundances on treated and control plots (univariate ANOVA, p <0.05, Table 5.5) and for s-methoprene, both Acariformes and Collembola showed different abundances. However, the Treatment x Time interaction was not significant in all cases, even after post- hoc pooling of the Plot (Treatment x Time) term. Furthermore, any differences between treated and control plots were generally inconsistent, especially between the two locations with more changes in abundance being recorded at Coomera Waters compared with Garden Island.

The Acariformes were composed primarily of members of the Family Oribatulidae (Sarcoptiformes: Oribatida) at both localities. The number of Acariformes per sample was low (range: 0 to 68 individuals) and variable (range of CV: 43 to 233%). There were three temporal trends in abundance that were evident: a decrease towards day 20, an increase over time or a peak in the middle of the 20 day period (at 5 or 8 days; Figure 5.2). Regardless of the temporal pattern, the abundance of Acariformes on treated and control plots tended to follow the same pattern; except for three occasions when the abundance of Acariformes showed an increase on treated plots (Table 5.5; Figure 5.3): after the Bti treatment at Coomera Waters (pitfall traps: p <0.0001), after the s-methoprene treatment at both Coomera Waters (aspirator: p = 0.023), and at Garden Island (aspirator: p = 0.007). The effect of the Bti treatment interacted with time (p = 0.022) and there were significantly more Acariformes in treated plots than the control plots on day 8 (post hoc: p = 0.002) but by day 20, abundances were similar, indicating that the response of Acariformes was short-term. For s-methoprene, the Treatment x Time interaction was not significant; however, it is evident from Figure 5.3 that the abundance of Acariformes was higher on day 5 at Coomera Waters and day 8 at Garden Island, but by 20 days, the abundance of Acariformes was similar to control levels at both localities. At these times, simultaneous samples were collected with either pitfall traps or aspirators; however, the increase in numbers was not detected with both techniques (Bti, Coomera Waters, aspirator: p = 0.477; s-methoprene, Coomera Waters, pitfall traps: p = 0.257; s-methoprene, Garden Island, pitfall traps: p = 0.078). Ecological impact assessment 145

Control Treatment

16 Bti (CW; A; p = 0.477) 12 Bti (CW; PF; p < 0.0001*) 14 10 12 8 10 8 6

6 4 4 2 2 0 0 01 58 10 15 20 051015201 5 8 20

35 8 S-meth (CW; A; p = 0.023*) S-meth (CW; PF; p = 0.257) 30 7 25 6 5 20 4 15 3 10 2 5 1 0 0 01 55 8 10 15 2020 01 5 8 10 15 20 Total number of Acariformes per sample

18 4 S-meth (GI; A; p = 0.007*) S-meth (GI; PF; p = 0.078) 16 3.5 14 3 12 2.5 10 2 8 1.5 6 4 1 2 0.5 0 0 01 58 10 15 20 051015201 5 8 20 Days post treatment

Figure 5.3: Mean (± SD) abundance of Acariformes from treated and control plots sampled after treated plots were exposed to either Bti or s-methoprene (Locations: CW = Coomera Waters; GI = Garden Island; Sampling technique: A = Aspirator n = 4; PF = Pitfall trap n = 8; * p <0.05; Natural log data are back transformed). 146

Table 5.5: Differences in abundance of individual taxa between treated (Bti and s- methoprene) and control plots, compared with univariate two-way analysis of variance (using STATISTICA).

p value

Taxa Treatment Localitya Sampleb Treatment Time Interaction

Total individuals Bti CW A 0.392 0.002 * 0.815 PF 0.848 0.027 * 0.192 SN 0.923 0.727 0.021 * GI A 0.990 0.001 * 0.965 PF 0.812 <0.001 * 0.916 SN 0.004 * 0.001 * 0.035 * s-meth CW A 0.924 0.023 * 0.436 PF 0.432 <0.001 * 0.328 SN 0.066 0.054 0.294 GI A 0.367 0.025 * 0.842 PF 0.658 0.001 * 0.965 SN 0.723 <0.001 * 0.031 *

Adult Diptera Bti CW A 0.598 0.275 0.842 GI A 0.826 0.003 * 0.981 PF 0.787 0.015 * 0.846 s-meth CW A 0.093 0.029 * 0.707 GI A 0.510 0.045 * 0.857

Heteroptera Bti CW A 0.051 0.081 0.765 PF 0.870 <0.001 * 0.002 * s-meth CW A 0.794 0.459 0.477 PF 0.366 <0.001 * 0.815 GI PF 0.721 0.029 * 0.162

Hymenoptera Bti CW A 0.792 <0.001 * 0.908 PF 0.690 0.011 * 0.318 s-meth CW PF 0.822 0.009 * 0.120 GI PF 0.955 0.044 * 0.954

Ecological impact assessment 147

Table 5.5 cont.

p value

Taxa Treatment Localitya Sampleb Treatment Time Interaction Collembola Bti CW PF 0.776 0.042 * 0.336 s-meth CW PF 0.028 * <0.001 * 0.524 GI PF 0.030 * 0.219 0.496

Acariformes Bti CW A 0.477 0.100 0.606 PF <0.001 * 0.175 0.022 * s-meth CW A 0.023 * <0.001 * 0.716 PF 0.257 <0.001 * 0.307 GI A 0.007 * 0.084 0.270 PF 0.078 0.785 0.086

Araneae Bti GI PF 0.828 <0.001 * 0.897

Copepoda Bti CW SN 0.901 0.716 0.021 * GI SN 0.003 * 0.003 * 0.057 s-meth CW SN 0.065 0.054 0.285 GI SN 0.748 <0.001 * 0.034 *

Grapsidae Bti GI PF 0.960 <0.001 * 0.914

Notes Data were transformed to natural log where necessary to meet the assumptions of homoscadasticity after Cochran’s test. * p <0.05. a. CW = Coomera Waters; GI = Garden Island. b. A = Aspirator; PF = Pitfall; SN = Aquatic sweep net.

148

The Collembola were composed primarily of Hypogastrura vernalis (Carl) (Hypogastruridae) at both localities. No consistent differences in Collembolan abundances between treated and control plots were recorded at the two localities after s-methoprene treatments. The abundance of Collembola increased on treated plots at Coomera Waters (pitfall traps; p = 0.028) but decreased at Garden Island (pitfall traps; p = 0.030; Table 5.5; Figure 5.4); furthermore, the Treatment x Time interaction was not significant at either locality (Coomera Waters: p = 0.524; Garden Island: p = 0.496). The effect of Time was significant at Coomera Waters reflecting the fluctuations in abundance within the monthly tidal cycle. Generally, collembolan abundance was low directly after the flooding tide and increased as the saltmarsh dried out. But after the Bti treatment at Coomera Waters, abundances remained low on both treated and control plots and no difference in the number of Collembola sampled with pitfall traps was recorded (p = 0.776). There was no general trend to suggest any consistent effect of Bti and s-methoprene treatment on collembolan abundance. This inconsistency may suggest that the dynamics of the abundance of Collembola differed among the localities. Although a significant decrease was recorded, this difference cannot easily be interpreted as an effect of treatment.

The Subclass Copepoda was composed of the Families Cyclopidae, Harpacticoida and Calanoida at both localities. There was considerable variation of the abundance of copepods among replicate samples (range: 6 to 6150 individuals; range of CV: 33 to 156%). However, a temporal trend was evident with numbers either increasing or decreasing with time (and this was significant at Garden Island, Table 5.5, Figure 5.5). The same temporal pattern was followed by treatment and control plots three out of four times: after the Bti treatment at Coomera Waters and both s-methoprene treatments. In contrast, after the Bti treatment at Garden Island, there were significantly fewer copepods in treated pools (p = 0.003) and the effect did not interact with time (p = 0.052). Ecological impact assessment 149

Control Treatment

6 Bti (CW; p = 0.776) 5

0 01 58 10 15 20

14 S-meth (CW; p = 0.028*) 12 10 8 6 4 2 0 051015201 5 8 20 Total number of Collembolan per sample

10 S-meth (GI; p = 0.030*) 9 8 7 6 5 4 3 2 1 0 01 55 8 10 15 20 Days post treatment

Figure 5.4: Mean (± SD) abundance of Collembola from treated and control plots sampled in pitfall traps after treated plots were exposed to either Bti or s-methoprene (Locations: CW = Coomera Waters; GI = Garden Island; n = 8 [two replicate samples from four plots]; * p <0.05; Natural log data are back transformed). 150

Control Treatment

1600 Bti (CW; p = 0.901) 1400 Bti (GI; p = 0.003*) 1400 1200 1200 1000 1000 800 800 600 600 400 400 200 200 0 0 012341 4 012341 4

600 250 S-meth (CW; p = 0.065) S-meth (GI; p = 0.748) 500 200 400 150 300 100 Estimated total number of Copepoda per200 sample

100 50

0 0 012341 4 012341 4 Days post treatment

Figure 5.5: Mean (± SD) abundance of Copepoda from treated and control plots after treated plots were exposed to either Bti or s-methoprene (Locations: CW = Coomera Waters; GI = Garden Island; n = 8 [two replicate samples from four plots]; *p <0.05; Natural log data are back transformed). Ecological impact assessment 151

Experimental applications of Bti and s-methoprene did not result in significant changes in the abundance of adult Diptera, Heteroptera and Hymenoptera relative to the control plots. Additionally, Bti did not impact on Araneae, Collembola and Grapsidae. These taxa were identified as contributing significantly to the composition of the communities using SIMPER analyses. The magnitude and direction of spatial and temporal changes of variability between treated plots were within the natural range shown by the control plots. Therefore, there was no evidence that Bti or s-methoprene changed the abundance of these taxa. 152

5.4 Discussion

The design of the experiment was intended to mimic the mosquito control treatments conducted in south-east Queensland and therefore provide a realistic indication of whether such treatments cause impacts on non-target macro-invertebrates. Generally, the results suggest that treatments of Bti and s-methoprene did not lead to a decrease in diversity or in the total abundance of arthropods. Some changes in the abundance of Acariformes, Collembola and Copepoda were recorded; however, the differences were not spatially and temporally consistent or in agreement with predictions. For example, the number of Acariformes appeared to increase significantly on treated plots. This was contrary to the predicted decrease in abundance of micro-invertebrates recorded by Hershey et al. (1998) in lentic freshwater. A decreased abundance of Copepoda was recorded after one of the Bti treatments. However, this change was not spatially consistent.

This study attempted to test the hypothesis that applications of Bti and/or s-methoprene to experimental plots would cause the assemblages of non-target arthropods to change in a way that was different from the untreated control plots. Previously, experimental manipulations at similar scales have proven to be useful for detecting disturbances in intertidal wetlands (Skilleter and Warren 2000, Skilleter et al. 2005). The testing of correlative patterns between an entire treated locality and reference localities can be used to identify environmental impacts (e.g. Underwood and Peterson 1988, Stark 1998). However, unambiguous evidence of causal relationships is best derived from manipulative experiments (Lindegarth and Underwood 2002) such as the current trial. Although weak changes in the composition and/or relative abundances of different taxa were recorded after the application of Bti and s-methoprene at Coomera Waters, these changes were not evident when the individual taxa were examined.

In this study, it was attempted to carry out the experimental manipulations at a spatial scale at which the response of non-target arthropods would be detected, i.e. entire ephemeral pools, and at a scale that was also practical. Although routine applications of Bti and s- methoprene are applied to the entire saltmarsh, there was no evidence of small-scale change that could be scaled up to the entire saltmarsh. In contrast, Hershey et al. (1998) reported a long-term reduction in insects in lentic freshwater wetlands in North America. However, the freshwater wetlands examined in her study are fundamentally different to the subtropical Ecological impact assessment 153 saltmarshes of south-east Queensland, in terms of hydrology and food web structure. This fundamental difference and the colonisation of different species could offer one explanation for the different responses of the communities to Bti and s-methoprene exposure.

Higher numbers of Acariformes (Family: Oribatulidae) were recorded on treated plots, at three times during the trial and were not related to location or treatment. The Oribatulidae are secondary decomposers that predominantly feed on litter material, microorganisms and fungi (Schneider et al. 2004). Considering this, it is difficult explain the short-term (5 – 8 day) increase in the Acariformes population on terrestrial plots. Similarly, the abundance of Collembola was affected by the application of s-methoprene on two occasions; however, the response was confounded by the location of the experiment. At Coomera Waters the abundance was higher on treated plots, while at Garden Island a lower abundance was recorded. The decreased abundance of copepods recorded after one of the Bti treatments was also spatially inconsistent. The abundances of Acariformes, Collembola and Copepoda from treated and control plots were extremely variable in space and time. On the terrestrial plots, the abundance of the Acariformes and Collembola had high temporal variability as the adults could readily move in and out of the plots. In the aquatic pools, the abundance of Copepoda had high spatial variability as swarms form during daylight hours (Ambler et al. 1991, Leising and Yen 1997).

Changes in abundance of invertebrates at the scale tested during the trial (hundreds of metres) are normal (Chapman et al. 1995, Barnes and Chapman 1998). The changes in abundance between treated and control plots that were recorded may reflect the heterogeneous nature of Acariformes, Collembola and Copepoda and the chance of getting large numbers in some replicates. The experimental design used here was specifically planned to allow detection of effects which varied in size among space and time. These data show that changes recorded on the treated plots was within the range of the controls. Therefore, although abundances of Acariformes, Collembola and Copepoda did change on some treated plots after spraying, these temporal changes could not be attributed to application of Bti or s-methoprene.

The results indicated that neither Bti nor s-methoprene impacted the abundance of adult Diptera, Heteroptera and Hymenoptera relative to the control plots. Additionally, Bti did not impact on Araneae, Collembola and Grapsidae. However, it is thought that s-methoprene 154 may have chronic impacts on organisms that undergo similar moult cycles to Diptera. The chronic effects of s-methoprene in the laboratory have been recorded to include: weight loss and delayed release of brood of estuarine mysid (Mysidopsis bahia: McKenney and Celestial 1996), altered shell formation of the blue crab (Callinectes sapidus: Horst and Walker 1999) and reduced growth and altered reproduction of branchiopods (Daphnia magna: Templeton and Laufer 1983, Olmstead and LeBlanc 2001, 2003). It is unknown if such chronic impacts would transfer into a field situation. If chronic impacts did occur, they could be detected as a change in abundance (Ward et al. 1976, Underwood 1995, Chapman 2002) of the specific taxa, which was not recorded during the current trial. However, it is possible that periods longer than 20 days may be required for such changes to be recorded.

Authors of previous studies from the northern hemisphere have indicated that s-methoprene was more broadly toxic to non-target organisms than Bti (Hershey et al. 1998). However, this was not true in Australia. The literature from the northern hemisphere indicates that high dosages of Bti applied for mosquito control may impact on non-target Chironomidae (Ali 1981, Car and de Moor 1984, Back et al. 1985, Hershey et al. 1998, Rey et al. 1998, Niemi et al. 1999), with minimal impacts on other insects (Hershey et al. 1995). In contrast high dosages of s-methoprene have been suggested to impact on a range of dipteran species (Miura and Takahashi 1973, Fargal and Temerak 1981) especially Chironomidae (Miura and Takahashi 1973, Ali 1991, Hershey et al. 1998), as well as other insects such as Ephemeroptera (Norland and Mulla 1975). Insufficient numbers of Chironomidae and Ephemeroptera were collected during the present study to examine the effects of Bti and s- methoprene application.

The results of the current trial concur with Barnes and Chapman (1998) who did not record an effect of Bti on the community composition of insect larvae (Chironomidae), crustaceans and molluscs (Arthritica helmsii and Xenostrobus securis) in saltmarshes in New South Wales. Their study used a before/after design, with adequate temporal and spatial replication. Barnes and Chapman (1998) focused on benthic organisms, which were collected in sediment samples, where the current study collected mobile terrestrial and aquatic arthropods. This suggests that Bti does not affect the entire range of organisms that inhabit saltmarshes. Ecological impact assessment 155

The multivariate analysis of variance did suggest a weak impact of Bti and s-methoprene treatment on the composition and/or relative abundances of different taxa; however, this was related to the location of the experiment. The impact on the community composition was minimal at Garden Island, which is a pristine island situated in Moreton Bay, with no development or obvious impacts. In contrast at Coomera Waters, where the intensity of the impact was more prominent, the location is bordered by farmland and canal estates on the Gold Coast. It is possible that at this location, the organisms may be already exposed to stress from runoff and the disturbed environment. Potentially, the differences in the quality of the environment may account for why these groups of animals responded differently. Where organisms are under pressure from other pollutants, the addition of Bti or s- methoprene may be enough to alter the ecosystem (Chapman 2002) and it is possible that other pollutants or the combination of pollutants are responsible for the observed changes. More detailed experimentation at Coomera Waters would be required to determine if the communities were responding to the presence of Bti and s-methoprene or other stressors in the ecosystem.

The locations also differed in topography and vegetation. The amplitude of the tidal flat was lower at Garden Island, meaning that the location was flooded more frequently compared with Coomera Waters. Also, at Coomera Waters the saltmarsh was primarily colonised by S. virginicus where Garden Island was primarily colonised by S. quinqueflora. The different vegetation of the locations may have influenced the relative abundance of organisms collected in pitfall traps (Melbourne 1999).

In conclusion, it was not possible to detect any consistent changes on the assemblages of non-target arthropods in the saltmarshes in subtropical Queensland. Based on the literature, both products appear safer than the organophosphate alternatives. The organophosphates registered for mosquito control are broad spectrum insecticides and are toxic to a range of Arthropoda, including Insecta (Porter and Goimerac 1969, Hurlbert et al. 1972, Didia et al. 1975) and Crustacea (Ludwig et al. 1968, Didia et al. 1975, Mortimer and Chapman 1995, Guzzella et al. 1997, Key et al. 1998, Appendix B). Furthermore, some organophosphates, e.g. chlorpyrifos, have been recorded to bioaccumulate through the food chain (Varo et al. 2002). 156

The results of this trial indicate that applications of either Bti or s-methoprene are one of the most environmentally sound methods of mosquito control currently available. Applications of both products did not decrease the diversity or abundance of non-target arthropods and both appeared to be of equal environmental desirability. The saltmarshes in south-east Queensland are being exposed to increasingly intense and more diverse combinations of human disturbance (Australian State of the Environment Committee 2001, Adam 2002). Efficient management of the ecological, ethical and economic values of saltmarshes depends on whether the disturbances which cause damage and those that do not can be identified. With such knowledge, informed decisions could be made about which disturbances must be removed and which, in fact, do not cause any impacts and can be tolerated (Underwood and Peterson 1988). The results of these trials suggest that applications of Bti and s-methoprene will not impact on the long-term structure and composition of arthropod assemblages in saltmarshes. This information is essential for developing the most environmentally sound mosquito control operations possible.

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Chapter 6

Concluding discussion

6.1 Assessing the use of new insecticides

During this thesis, a practical and repeatable process was followed to assess the potential of new insecticide formulations for use by a control industry, whether it is public health, agriculture or domestic purposes (Figure 6.1). Firstly, laboratory studies were used to provide base-line information, followed by field trials to assess efficacy and application equipment. Finally, the environmental impacts were assessed.

The initial laboratory trials provide a quick and economical means to gather data on the potential of a product for control. The methodologies for assessing the susceptibility of a target organism to insecticides have been outlined by the World Health Organisation (WHO 2005). However, it can be difficult to extrapolate results into the field where the environment and climate may alter the efficacy of a product (Chapter 2).

The most accurate method to assess the efficacy of a product in the field is to monitor the mortality of caged organisms, which is a quick, simple and economical method. A known number of individuals are exposed to the treatment in replicated mesh cages, and the surviving individuals are re-counted after a specified time. In contrast, the accuracy of sampling naturally occurring populations is highly variable and statistical comparison of trials conducted at different times was problematic (Chapter 3).

Before a new formulation can be adopted into control programs, the effectiveness of broad- scale applications must be assessed, to ensure that the correct amount of insecticide can be applied in a uniform manner to the target area. Due to the expense and logistics involved in conducting such trials, it is imperative that trials are well designed and that the size of catch- 158

New insecticide formulation

No Are the common species susceptible in the laboratory?

Yes

No Are the major target species susceptible in field situations?

Yes

No Can the product be adapted into broad-scale control operations?

Yes

No Are the environmental impacts acceptable?

Yes

Incorporate into control operations

Public Habitat Biological Early detection education modification control systems

Figure 6.1: Proposed process for investigating the possibility of incorporating new insecticide formulations into current control operations. Concluding discussion 159 trays used is appropriate. The results in Chapter 4 showed that 1 m2 catch trays were too small to sample products with a large granule size, such as VectoBac G and Altosid pellets. It is recommended that catch trays are at least 2 m2 to sample VectoBac G and 4 m2 to sample Altosid pellets. The number of catch trays, required to estimate the mean with 95% accuracy, can be selected from Table 4.3.

To examine the ecological impacts of insecticide application, there is clearly a need for field-based trials. However, the populations of non-target animals which need to be sampled are dynamic and undergo natural fluctuations. Sampling must be sufficient to identify unusual patterns of change in interactive and variable measurements. Thus, it is important that the accuracy and power of sampling tools are optimised for each experiment. During the pilot trial in Chapter 5, there was a large disparity in the number of organisms among replicates collected with the initial sampling techniques. As a result, any changes in the abundance and composition of non-target communities may have been concealed, as the power to detect any differences would have been relatively low. Using information from pilot trials, the sampling gear can be altered to accurately sample during sequential trials to provide a realistic indication of the impact of insecticide applications. Although variations in the abundance of saltmarsh invertebrates is normal, the design of the manipulative experiment followed in Chapter 5 provided a powerful tool for assessing the causal relationship of an environmental impact.

6.2 Contribution to mosquito control operations

To date, effective broad-scale control of ground-pool freshwater larvae has proved difficult to achieve. Bti is the main product used to control ground-pool freshwater species, yet the efficacy various formulations had not previously been defined in the Australian environment. This study determined that Bti (VectoBac 12AS and VectoBac WG) was effective for the control of freshwater ground-pool mosquito larvae and can confidently be incorporated into control programs. The products lost little efficacy in the field (Chapter 3) compared with the laboratory-based data (VectoBac WG: Chapter 2; VectoBac 12AS: Table 2.2). The laboratory trials also indicated that pyriproxyfen may be useful for the control of 160

Cx. annulirostris in freshwater ground-pools. However, field trials need to be conducted with this product.

To control larvae in container habitats, operators rely largely on community participation. At present, the only product available for domestic use is granular s-methoprene available at hardware stores. Yet, Bti is seen as a more desirable product for community use, due to the quick acting nature of the endotoxins. The results of the laboratory trials in Chapter 2 indicated that Bti (VectoBac WG) was effective for the control of container inhabiting larvae, and was active in the field against Ae. notoscriptus for up 1 week. In contrast, pyriproxyfen was unlikely to be useful for control in container habitats; it was not effective against Ae. aegypti and Ae. notoscriptus in the laboratory.

Mosquito control in estuaries is well developed and broad-scale control is regularly carried out, but the treatment of heavily vegetated areas is still more problematic than in open areas. When treating heavily vegetated areas, granular formulations are useful to penetrate the foliage. However, there are no economical granular formulations available. Granular s- methoprene (Altosid pellets) is sometimes used, but it is expensive at AUD $335 per ha (applied with a Bell 47 helicopter); furthermore, this product is extremely difficult to apply evenly to the target site (Chapter 4). Although the granular Bti formulation (VectoBac G) is cheaper to apply than granular s-methoprene, there are difficulties associated with broad- scale application.

Granular Bti is bulky and has a low potency meaning that large amounts of product must be carried to treat a small area. It was found in Chapter 3 that VectoBac G is likely to start failing below 266 Bti ITU/L (2 kg/ha) against Ae. vigilax in saltmarsh pools. Considering this, a target application rate for aerial applicators was identified as 5 kg/ha (or 666 Bti ITU/L in 15 cm deep water) for use in Australian conditions, while still providing a sufficient safety margin for application error, in terms of control. Applying product at this rate would reduce operating costs by almost 30% (price reduced from AUD $63/ha for 7 kg/ha to $45/ha for 5 kg/ha applied with a Bell 47 helicopter). During the aerial trial (Chapter 4), it was difficult to distribute VectoBac G evenly at 5 kg/ha; despite this, the treatment was successful in terms of control and there was 100% mortality of the target larvae. The most important factors that affected the observed spatial distribution of product were the uneven flight path of the helicopter and the low sensitivity of the catch trays. The Concluding discussion 161 results of Chapter 4 provide useful information to local government operators about a product that is rapidly increasing in use; over 12,000 kg of VectoBac G have been applied per year in south-east Queensland since 2003 (Figure 1.1).

Applications of Bti and s-methoprene did not decrease the diversity or abundance of non- target arthropods in south-east Queensland. Some changes in the abundance of Acariformes, Collembola and Copepoda were recorded; however, the differences were not spatially and temporally consistent or in agreement with predictions. For example, after application of Bti to ephemeral pools, Copepoda were sampled in lower numbers at only one location and no differences were recorded after treatments of s-methoprene to ephemeral pools. After applications of s-methoprene to terrestrial plots, higher numbers of Acariformes were recorded at both localities, and this was also recorded after application of Bti at Coomera. Any changes in the abundance of taxa were recorded within 8 days of the experimental treatments and abundances had converged with control levels by 20 days. The changes in abundance of arthropods between treated and control plots that were recorded may reflect the large spatial variability of arthropods and the chance of getting large numbers in some replicates.

Authors of previous studies from the northern hemisphere have indicated that s-methoprene is more broadly toxic to non-target organisms than Bti and this has shaped local management policy in terms of the product choice. My results highlight that generalising from results in the northern hemisphere is inappropriate for Australia. Compared to the literature concerning the environmental impacts of organophosphates, both products appear safer than temephos and chlorpyrifos. Overall, the effects of Bti and s-methoprene were minimal and unlikely to have long-term environmental consequences.

These results provide essential information to local government operators who are continually taking steps to advance current mosquito control operations. By refining the world-class mosquito control operations in Queensland with the information provided in this thesis, local governments can take steps towards developing a safe and healthy environment for use by the rapidly growing population. 162

6.3 Insecticides as part of an integrated pest management plan for mosquito control

Successful control programs integrate different control options. The different mosquito control strategies include habitat modification, biological control, public education and insecticide use. Control options should be conducted on a case-by-case basis, after the efficacy and environmental impacts of each option are considered. Recently a statistical method for identifying outbreaks of Ross River virus was developed and has formed the basis of a web-based early warning system for Queensland, identifying times when rapid mosquito control by local governments is required (Gatton et al. 2004).

Physical habitat modification (e.g. runnelling) involves digging channels through wetland areas, such as saltmarsh, to link isolated pools with the tidal source. This increases water movement during low amplitude tides, providing for more regular flushing of stagnant water (Hulsman et al. 1989). Consequently, larval development (Hulsman et al. 1989) and egg conditioning (Dale et al. 2002) of mosquitoes is unsuccessful. Runnelling also increases access for larvivorous fish to forage in the saltmarsh (Hulsman et al. 1989). Runnelling is relatively cheap to implement, however to be effective, the runnels must be maintained by local councils (Hulsman et al. 1989). Physical habitat modification has been used extensively in the USA (Ferrigno and Jobbins 1968, Shisler and Jobbins 1977) and parts of Australia (Easton 1986, Hulsman et al. 1989). The decision whether to modify a site or to apply insecticides may be determined by the physical properties of the site, as some wetlands are not amendable to runnelling (Russell and Kay 2004) and the application of insecticides remains the most common approach in most parts of the world (Federici 1995, Russell and Kay 2004).

Runnelling has been recorded to increase the transportation of mangrove propagules into the saltmarsh (Breitfuss et al. 2003) and has been demonstrated to affect the abundance of gastropods, crabs, prawns and fish because it imposes a lower datum; however, changes are isolated to within 10 m of runnels (Dale and Dale 2002, Breitfuss 2003, Breitfuss et al. 2004, Connolly 2005). Although runnelling may lead to isolated community changes on a small area of the saltmarsh, it has been demonstrated to have minimal long-term impacts on Concluding discussion 163 the entire saltmarsh (Dale and Dale 2002, Jones et al. 2004). This is comparable with the application of Bti and s-methoprene.

Biological control agents (e.g. fish and copepods) are successful in simple and confined ecosystems, such as wells and containers (Hurst 2004, Kay and Nam 2005). A successful example is the control of Ae. aegypti in water storage containers in Vietnam using copepods (Mesocyclops spp.; Nam et al. 1998, 2000, Kay and Nam 2005). Biological control is situation dependant and, although successful in Vietnam, the use of copepods for the broad- scale control of Ae. aegypti in Australia is not likely to be as successful. Copepods are not suitable for application in small containers with a high turnover of water (i.e. discards and roof gutters) which are key containers during the summer (Montgomery and Ritchie 2002); however, copepods were effective in subterranean habitats (Russell et al. 1996a), which provide key containers during the winter (Russell et al. 1997). In some situations, biological control alone may not be adequate to control naturally occurring mosquito populations and may be used in conjunction with insecticides (Riviere et al. 1987, Tietze et al. 1994, Hurst 2004) thus, the use of environmentally sound insecticides will still be required.

Biological control in artificial environments with indigenous organisms is unlikely to result in any substantial environmental impacts. Although in the case of fish, the water quality for drinking may be reduced by defaecation and excretion of oils (Fletcher et al. 1992). If non- native fish are introduced into large water bodies (such as freshwater ground-pools) for control purposes, significant environmental impacts may occur. This has occurred in Australia with the introduction of Gambusia affinis holbrooki (Baird and Girard) in the 1920s, which is now widespread and is thought to have played a significant role in the decline of many native freshwater fish (Arthington et al. 1986, Lloyd et al. 1986, Arthington and Marshall 1999). Hurst (2004) has recommended the use of native freshwater fish, Melanotaenia duboulayi (Castelnau) and Hypseleotris galii (Ogilby), as alternatives to G. holbrooki; however, M. duboulayi should be introduced into water bodies with caution as it is a voracious predator of insects and tadpole populations (Hurst 2004).

164

6.4 Mosquito control in perspective

Insecticides are used to control pests in a range of industries which broadly includes public health, domestic and agricultural purposes. The primary insecticides used for agricultural purposes are a range of organophosphate based products, including parathion methyl, chlorpyrifos, dimethoate, profenfos and diazinon (Radcliffe 2002). Approximately 50,000 tonnes of organophosphates are used by the agricultural industry each year in Australia (Radcliffe 2002), compared with less than 200 kg for mosquito control (Figure 2.1, CLAG 2005, NEMMO 2005).

The organophosphates used by the agricultural industry tend to be extremely effective for the control of a range of pests. However, run-off has resulted in severe environmental impacts and residues have been detected in biota and waterways near intensive agricultural areas (Barron and Woodburn 1995, Hamilton and Haydon 1996, Russell et al. 1996b, Müller et al. 2000). As awareness increases of the environmental impacts of organophosphate use, farmers are developing integrative pest management plans, which include the use of biologically-based insecticides and transgenic plants (Radcliffe 2002). The move towards integrative pest management and environmentally “softer” insecticides has not been adapted by the agricultural industry as rapidly as in the mosquito control industry.

To put this into perspective, recent changes in insecticide use by the cotton industry have been made due to increasing environmental concerns, including toxicity to non-target organisms (Hurlbert et al. 1972, Barron and Woodburn 1995), runoff into streams and estuaries (Russell et al. 1996b, Mortimer 2000) and contamination in beef (Woods et al. 2001a). This lead to the use of transgenic plants that have a single Bacillus thuringiensis (Bt) endotoxin incorporated into the deoxyribonucleic acid (DNA), and are toxic to pest lepidopteran larvae when ingested (Luttrell et al. 1994). Approximately 30% of Australia’s cotton crop is currently sown with Bt cotton and on these areas insecticide use has been reduced by half, resulting in an overall reduction in organophosphate use across the industry of 12 – 15% (Radcliffe 2002). However, as only one endotoxin was incorporated into the plants, the lepidopteran larvae quickly developed resistance (Tabashnik 1994, Gould 1998). In contrast, resistance of mosquitoes to Bti has not been recorded in the field and this can be contributed to the use of the entire complex of endotoxins (Georghiou and Wirth 1997). Concluding discussion 165

Furthermore, the use of Bti for mosquito control has nearly completely replaced organophosphate use.

Insect growth regulators, including diflubenzuron, triflumuron, fenoxycarb, s-methoprene and tebufenozide, have been incorporated into agricultural practices in recent years, for the control of invertebrate pests including the flour beetle, grain beetle and weevils (Radcliffe 2002). However, this has done little to curtail organophosphate use, as the use of insect growth regulators has been limited by their slow action and the narrow range of sensitive stages in the pest life cycle (Casida and Quistad 1998). For each formulation, about 1 tonne is used per year in Australia (Radcliffe 2002). This is comparable to mosquito control, which applies about 1 tonne of s-methoprene per season (Figure 2.1), but trivial compared with the amount of organophosphates used by the agricultural industry.

6.5 Conclusions

Where organised and broad-scale mosquito control programs have been implemented in Australia, a reduction in the incidence of mosquito-borne disease, mainly Ross River virus, has been recorded (Gatton et al. 2004). In spite of this, over 77,000 human infections have been recorded since 1991 (Communicable Diseases Network – Australia New Zealand – National Notifiable Diseases Surveillance System). There remains a high incidence of Ross River and Barmah Forest viruses among working adults, which is of considerable public health concern. Currently the social and economic costs of mosquito-borne disease are unclear; however, each case of Ross River and Barmah Forest virus is estimated to cost between AUD $590 – $2,500, equating to an estimated total cost of AUD $5.6 – over $12.5 million per year, with over 50% of these costs incurred by the state of Queensland (Russell 1998b, Harley et al. 2001). Similarly, there are no precise estimates of the national cost of mosquito control and surveillance programs, but based on individual budgets the expenditure is thought to be in the order of AUD $15 – $20 million per year.

There is also a concern that the risk of disease transmission will increase in the future. This is likely to result from a downgrading of vector control activities, lack of political will, 166 insufficient funding, rapid international travel, growing urbanisation with insufficient infrastructure (i.e. water supply and sanitation), poverty and global warming (Russell 1998b, Transer et al. 2003, Russell and Kay 2004). As a result, continual progress will need to be made in mosquito control practices in order to remain proactive.

The results of this thesis demonstrate that Bti can be used to effectively control mosquito immatures under different field conditions; and that Bti and s-methoprene did not cause an adverse impact on invertebrate communities in saltmarshes. Considering that the appropriate use of Bti and s-methoprene can reduce the incidence of arbovirus transmission among the local human population, the future application of these products is supported. The use of insecticides should be integrated with public education, biological control, physical habitat modification and early detection systems.

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213

Appendix A

Publications relevant to thesis

Russell, T. L., M. D. Brown, D. Purdie, P. A. Ryan, and B. H. Kay. 2003. Efficacy of VectoBac (Bacillus thuringiensis var. israelensis) formulations for mosquito control in Australia. Journal of Economic Entomology 96: 1786-1791.

Russell, T. L., and B. H. Kay. 2005. The use of Bacillus thuringiensis var. israelensis for the control of arbovirus vectors in the Australian environment. Arbovirus Research in Australia 9: 337-343

Russell, T. L., Kay, B. H. and Ryan, P. A. 2005. Mosquito control and Bacillus thuringiensis var. israelensis use, an Australian perspective, pp. 57-72. In N. D. Binh, R. J. Akhurst and D. Dean [eds.], 5th Pacific Rim conference on the biotechnology of Bacillus thuringiensis, Hanoi Vietnam.

ECOTOXICOLOGY Efﬁcacy of VectoBac (Bacillus thuringiensis variety israelensis) Formulations for Mosquito Control in Australia

TANYA L. RUSSELL, MICHAEL D. BROWN,1 DAVID M. PURDIE, PETER A. RYAN, AND BRIAN H. KAY

Queensland Institute of Medical Research and University of Queensland Australian Center for International and Tropical Health and Nutrition, P.O. Royal Brisbane Hospital, Brisbane, Queensland 4029, Australia

J. Econ. Entomol. 96(6): 1786Ð1791 (2003) ABSTRACT Laboratory bioassays were conducted on the efÞcacy of a water-dispersible granule (WG) formulation of Bacillus thuringiensis variety israelensis (VectoBac WG; active ingredient [AI]: 3,000 Bti international toxic units [ITU]/mg) against third instars of six common Australian mosquito species, Aedes aegypti (L.), Ochlerotatus vigilax (Skuse), Ochlerotatus notoscriptus (Skuse), Culex sitiens Wiedemann, Culex annulirostris Skuse, and Culex quinquefasciatus Say. The normal model for

log-linear mortality data was used to determine laboratory 48-h LC50 and LC95 values. The target mosquito species tested were extremely sensitive to the VectoBac WG formulation, with the most

sensitive species (Cx. annulirostris and Cx. quinquefasciatus,LC95 value of 0.019 ppm) being twice as susceptible as the most tolerant (Oc. notoscriptus,LC95 value of 0.037 ppm). Cx. annulirostris was selected as a target species for a small-plot evaluation of VectoBac WG and VectoBac 12 aqueous solution (AS) ([AI]: 1,200 Bti ITU/mg) efÞcacy over time, in freshwater in southeastern Queensland, Australia. Replicated cohorts of caged third instars were exposed weekly to six concentrations of WG formulation (0.004Ð0.13 ppm) and three concentrations of the 12AS formulation (0.04Ð0.13 ppm). In water with high organic content, treatment concentrations of 0.008 ppm WG and 0.04 ppm 12AS and above produced signiÞcant larval control (Ն96%) at 48 h posttreatment, with no residual control at week 1. Water quality was not affected by treatment with either formulation.

KEY WORDS Aedes, Culex, Ochlerotatus, Bacillus thuringiensis variety israelensis, biocontrol

AFTER INITIAL DESCRIPTION BY Goldberg and Margalit The potency of Bti, an aerobic, Gram positive bac- (1977), the microbial insecticide Bacillus thuringiensis terial pathogen, against mosquito larvae is attributed variety israelensis (Bti) de Barjac has been integrated to four crystallized protoxins, which after ingestion are into programs designed to control mosquito vectors of dissolved and activated under the alkaline action of arboviruses, such as Ross River, Barmah Forest, and the midgut and intestinal proteinases. The released Dengue. Species incriminated with the transmission of toxins bind to receptors on the epithelial midgut cells, these arboviruses are from the Aedes, Ochlerotatus, leading to swelling and lysis of the affected cells and and Culex genera (Kay 1982; Kay et al. 1984; Watson eventually cause insect death (Georghiou and Wirth and Kay 1998, 1999; Boyd and Kay 1999, 2000; Ryan et 1997, Regis et al. 2001). al. 2000). Arboviruses cause a signiÞcant impact on Valent BioSciences have developed a water-dis- Australian public health (Russell 1995, 1998), and in persible granule (WG) formulation of Bti (VectoBac the absence of vaccines, the community is reliant on WG; active ingredient [[AI]]: 3,000 Bti international insecticide use to control mosquito vectors, as stipu- toxic units [ITU]/mg) for broad-scale application lated by the Health Regulations 1996, part 8. The against Australian mosquito vectors. With this prod- implementation of the Environmental Protection Act uct, the Bti toxins are formulated onto micropellets 1994, places a duty on local governments to operate that disperse rapidly when placed in water (Abbott mosquito control programs in a manner that mini- 1992). In 2000, the National Registration Authority mizes environmental harm. Growing environmental (now the Australian Pesticides and Veterinary Med- concerns over the use of organophosphorus larvicides icines Authority) registered VectoBac WG for use has lead to an increased use of Bti, which is considered against mosquitoes in fresh, brackish, and saltwater to be more environmentally sound (Hershey et al. habitats in Australia. Preliminary Þeld trials found 1995; Brown et al. 1998a, 1999, 2002). VectoBac WG to be as effective as the widely applied liquid VectoBac 12AS ([AI]: 1,200 Bti ITU/mg) for- 1 Current address: FMC Chemicals, P.O. Box 329, Hamilton Central, mulation for application against Ochlerotatus vigilax Queensland 4007, Australia. (Skuse) in saltmarsh habitats (Muller et al. 2001). In

0022-0493/03/1786Ð1791$04.00/0 ᭧ 2003 Entomological Society of America December 2003 RUSSELL ET AL.: EFFICACY OF Bti FORMULATIONS AGAINST AUSTRALIAN MOSQUITOES 1787 comparison with the liquid (AS, aqueous solution) Field Bioassays. A small-plot Þeld trial was con- formulation, the WG formulation has additional ad- ducted between February and June 2002 in Logan vantages of economical transport, storage, and longer City, southeastern Queensland to assess the Þeld ef- shelf life. Þcacy of VectoBac WG and 12AS against Cx. annu- Accordingly, this project was designed to assess lirostris, using methods similar to Brown et al. (2004). efÞcacy of the new VectoBac WG product for use by Replicates of caged larvae were exposed to six con- the Australian mosquito control industry. Before the centrations of the WG formulation (0.004, 0.008, 0.016, Þeld trial, laboratory concentration-response data for 0.031, 0.063, and 0.13 ppm) and three concentrations VectoBac WG against common Australian mosquito of the 12AS formulation (0.04, 0.08, and 0.16 ppm), in species was developed. Sequentially, the efÞcacy of independent pools. For each concentration, Þve treat- varying application rates of VectoBac WG and Vec- ment and three untreated control pools (1 m in di- toBac 12AS against Culex annulirostris Skuse were ameter, 0.8 m in depth) were selected, with one open- evaluated in the Þeld, and abiotic water variables were topped, 250-␮m cylindrical cage (0.5 m in depth by monitored to assess possible effects as a consequence 0.5 m in diameter) placed in each pool. Larvae were of insecticide applications. These results provide base- transported to the site as early second instars and line data for the development of cost-effective Þeld released into a cage placed in a holding pool 24 h applications and resistance management. before treatment. For assessment of control efÞcacy, 100 larvae were placed in each treatment and untreated control cage, Ϸ2 h before treatment. Materials and Methods Treatment concentrations were formulated by add- Mosquito Test Species. Toxicity studies were con- ing the appropriate weight of WG (weighed in the ducted for third instars of Oc. vigilax (Victoria Point laboratory on the morning of treatment) or volume of 2000), Culex sitiens Wiedemann (Coomera Island 12AS (measured with a pipette) to 5 liters of sieved 1996), Cx. annulirostris (Boondall 1997), Culex quin- habitat water. This was then stirred with a glass rod quefasciatus Say (Gold Coast 2001), Oc. notoscriptus until the formulation was homogeneous and then (Skuse) (Brisbane 1995), and two strains of Aedes poured evenly across the surface of the pool. The aegypti (L.) (Torres Strait 2000; Townsville 2000). numbers surviving at 48 h postapplication were Specimens were derived from colonies maintained in counted, and the percentage of mortality was calcu- the Queensland Institute of Medical Research insec- lated. To test duration of control, weekly additions of tary, using methods based on those described for Cx. 100 larvae were made to each treated and untreated annulirostris by McDonald et al. (1977) and for Oc. control bioassay cage. As with the initial cohort, mor- notoscriptus by Watson et al. (2000). tality was assessed at 48 h postintroduction. Weekly Laboratory Bioassays. Concentration-response data larval additions occurred until no statistically signiÞ- were developed for VectoBac WG against the six tar- cant difference in mortality was recorded between get species, detailed above, using standard laboratory untreated control and treatments. bioassay methods for testing larval susceptibility Field Abiotic Water Characteristics. A Horiba (WHO 1981, Brown et al. 2000). Third instars were (Kyoto, Japan) portable Þeld laboratory was used to exposed to serial doses of WG mixed in tap water, measure the abiotic water characteristics (pH, tem- which had been sitting for at least 24 h. The salinity of perature, salinity, dissolved oxygen, and turbidity [ne- the test water was 0% for the freshwater species (Ae. pholometric turbity units; NTUs]). Measurements aegypti, Oc. notoscriptus, Cx. annulirostris, and Cx. were taken in all treatment and untreated control quinquefasciatus) and 1% for the saltmarsh species pools immediately before treatment, at 48 h post treat- (Oc. vigilax and Cx. sitiens). Five replicates each of 20 ment, and then at weekly intervals until completion of third instars were introduced into glass beakers con- the Þeld assessment. taining 200 ml of test concentration. Test specimens Statistical Methods. The laboratory bioassay data were individually removed from holding trays and were analyzed using the normal model for log-linear distributed randomly among the test beakers. Five mortality (Finney 1971, Piegorsch and Bailer 1997; untreated control beakers holding 20 test larvae each PROC PROBIT, SAS Institute 2001). This method was in 200 ml of water without WG were used for each more appropriate than standard probit analysis, be- concentration. cause the data were not normally distributed. Zero Initially, a number of range-Þnding tests with counts were analyzed as concentrations of 0.000001 widely spread exposure concentrations (0.002, 0.017, ppm, to avoid inÞnite logarithmically transformed val- and 0.171 ppm) were conducted. Based on these tests, ues. This method was adopted in favor of AbbottÕs a narrow range of concentrations that straddled the formula (Abbott 1925) because it does not modify the effective range were then used to determine LC50 and exposure variable and thus has negligible impact on LC95 values, expressed as both ITU per liter and the the curve. EfÞcacy of the Þeld bioassays was analyzed concentration of product in the mixture. The numbers using a two-way analysis of variance (ANOVA), with surviving were counted at 48 h. Death or the lack of factors for concentration rate and time. To normalize reaction to gentle prodding with a glass pipette was the data, the proportion of mortality in the Þeld bio- the measured deleterious response. The test larvae assays was arc sine transformed (Anscombe 1948). were not fed during testing to minimize variability The possible effects of treatments on abiotic water caused by nutritional and metabolic condition. characteristics over time were analyzed using a one- 1788 JOURNAL OF ECONOMIC ENTOMOLOGY Vol. 96, no. 6

Table 1. Laboratory bioassay results of VectoBac WG exposure against six Australian mosquito species at 48 h

a ␹2 Ϯ Species LC50 (95% CL) LC95 (95% CL) (df) Slope SE Cx. sitiens (Coomera Island) 0.019 (0.018 Ͻ LC Ͻ 0.021) 0.031 (0.023 Ͻ LC Ͻ 0.036) 280 (43) 6.5 Ϯ 0.7 Cx. quinquefasciatus (Gold Coast) 0.005 (0.003 Ͻ LC Ͻ 0.006) 0.019 (0.015 Ͻ LC Ͻ 0.036) 158 (68) 2.3 Ϯ 0.2 Cx. annulirostris (Boondal) 0.004 (0.003 Ͻ LC Ͻ 0.005) 0.019 (0.015 Ͻ LC Ͻ 0.036) 22 (57) 4.6 Ϯ 0.4 Oc. vigilax (Victoria Point) 0.013 (0.012 Ͻ LC Ͻ 0.014) 0.021 (0.019 Ͻ LC Ͻ 0.024) 113 (43) 6.5 Ϯ 0.8 Oc. notoscriptus (Brisbane) 0.015 (0.013 Ͻ LC Ͻ 0.017) 0.037 (0.032 Ͻ LC Ͻ 0.046) 132 (53) 3.6 Ϯ 0.4 Ae. aegypti (Townsville) 0.017 (0.015 Ͻ LC Ͻ 0.018) 0.031 (0.023 Ͻ LC Ͻ 0.037) 199 (58) 5.9 Ϯ 0.7 Ae. aegypti (Torres Strait) 0.018 (0.016 Ͻ LC Ͻ 0.019) 0.026 (0.024 Ͻ LC Ͻ 0.029) 280 (98) 0.3 Ϯ 0.03

CL, conÞdence limits. a Concentrations in ppm. way ANOVA for each treatment rate. Where neces- a statistically signiÞcant difference between the efÞ- sary, an all-pairwise comparison test at P Ͻ 0.05 cacy of 12AS treatments at the different time periods ϭ Ͻ (Tukey 1953; cited in Zar 1999) was used to determine (F1,34 673.64; P 0.001) and for the varying con- ϭ signiÞcant differences within each factor. centration rates, including untreated controls (F3,34 89.31; P Ͻ 0.001). At 1 wk posttreatment, poor control was observed with mortality ranging between of 6.8 Ϯ Results 6.5 and 19.6 Ϯ 8.4%. Untreated control mortality was Laboratory Bioassays. After 48-h exposure to Vec- 9.0 Ϯ 3.7 and 11.8 Ϯ 8.7% at 48 h and 1 wk posttreat- toBac WG in laboratory bioassays, third instars of the ment, respectively, differing signiÞcantly from all con- freshwater species, Cx. annulirostris and Cx. quinque- centration rates at 48 h (P Ͻ 0.05), but not at week 1 fasciatus, were found to be the most sensitive, with (P Ͼ 0.05).

LC50 values of 12 ITU/liter (0.004 ppm) and 15 ITU/ Field Abiotic Water Characteristics. The water con- liter (0.005 ppm), respectively, and identical LC95 ditions during this study were characterized by high values of 57 ITU/liter (0.019 ppm) (Table 1). The turbidity (range 33Ð525 NTUs), warm water (range saltmarsh species, Oc. vigilax, was slightly more tol- 15.8Ð27.1ЊC), lowÐmedium oxygen content (range erant with LC50 and LC95 values of 39 ITU/liter (0.013 ppm) and 63 ITU/liter (0.021 ppm), respectively. Cx. sitiens, a cohabitant of saltmarsh pools, recorded the highest LC50 value of 57 ITU/liter (0.019 ppm) and an LC95 value of 93 ITU/liter (0.031 ppm). For the container-breeding species Ae. aegypti, LC50 and LC95 values of 54 ITU/liter (0.018 ppm) and 78 ITU/liter (0.026 ppm) were estimated, respectively, for the Torres Strait strain and 51 ITU/liter (0.017 ppm) and 93 ITU/liter (0.031 ppm), respectively, for the Towns- ville strain. Oc. notoscriptus had an LC50 value of 45 ITU/liter (0.015 ppm) and the highest LC95 value of 111 ITU/liter (0.037 ppm). Field Bioassays. Mortality of third-instar Cx. annulirostris recorded at 48 h posttreatment ranged from 85 Ϯ 3.6 to 100.0 Ϯ 0% in pools treated with VectoBac WG at rates between 0.004 and 0.13 ppm (Fig. 1). At this time, efÞcacy of the 0.004 ppm WG concentration rate was signiÞcantly lower than the 0.016, 0.063, and 0.13 ppm treatments (P Ͻ 0.05). The efÞcacy of WG treatments were signiÞcantly different between time ϭ Ͻ periods (F1,66 957.19; P 0.001) and the varying concentration rates, including untreated controls ϭ Ͻ (F6,66 34.40; P 0.001). One week after treatment, poor control was observed and mortality ranged between 4.0 Ϯ 6.9 and 14.2 Ϯ 12.6%. Untreated control mortality was 9.3 Ϯ 4.3 and 12.2 Ϯ 9.7% at 48 h and 1 wk posttreatment, respectively, differing signiÞcantly from all concentration rates at 48 h (P Ͻ 0.05), but not at week 1 (P Ͼ 0.05). With respect to the VectoBac 12AS evaluations, the mortality of third-instar Cx. annulirostris ranged be- Fig. 1. Field evaluation of VectoBac WG (3,000 Bti ITU/ tween 99.0 Ϯ 0.7 and 100.0 Ϯ 0% at 48 h posttreatment mg) and VectoBac 12AS (1,200 Bti ITU/mg) efÞcacy against at rates between 0.04 and 0.16 ppm (Fig. 1). There was Cx. annulirostris. December 2003 RUSSELL ET AL.: EFFICACY OF Bti FORMULATIONS AGAINST AUSTRALIAN MOSQUITOES 1789

1.33Ð10.02 g/liter), low salinity (range 0Ð0.01%), and rate for mosquito habitat, in particular for freshwater neutral pH(range 6.40Ð7.77). No statistically signiÞ- areas, the depth of the water should be considered. cant effect on abiotic water characteristics over time In line with other data, the activity of Bti was lost was recorded as a consequence of either WG or 12AS within 1 wk, and residual activity was not increased treatments. with higher concentrations of product (Karch et al. 1991, Gelernter and Schwab 1993, Kroeger et al. 1995, Fillinger et al. 2003). In our study, the test pools also had high organic/pollution loads, intense sunlight, and Discussion high larval densities outside of the bioassay rings. Such Bti-based VectoBac products were shown to be factors have previously been demonstrated to reduce highly effective against a range of common Australian the effectiveness of Bti products (Mulla et al. 1990, mosquito vectors. Under laboratory conditions, all Pusztai et al. 1991, Becker et al. 1992, Yousten et al. mosquito species tested were extremely sensitive to 1992, Glare and OÕCallaghan 2000) and despite these the VectoBac WG formulation, with the most sensitive adverse test conditions, both formulations were highly species (Cx. annulirostris and Cx. quinquefasciatus) effective. The current study also conÞrms that Bti being twice as susceptible as the most tolerant (Oc. products are benign in terms of abiotic water quality. notoscriptus), indicated by LC95 values. In contrast, Based on these results, the Bti-based microbial in- greater variation in susceptibility to VectoBac 12AS secticides seem to be an excellent choice for treatment was recorded by Brown et al. (1998b, 2001) under of freshwater larval habitats and loses little efÞcacy in laboratory conditions, Ae. aegypti (LC95 values con- the Þeld. Additionally, in the laboratory a range of verted to Bti ITU/liter, 886.46) was indicated to be at saltwater, freshwater, and container-breeding larvae least 36 times more tolerant of Bti than Cx. sitiens were shown to be highly susceptible to VectoBac WG.

(LC95 values converted to Bti ITU/liter, 24.37). In Finally, a cost comparison of the minimum WG and terms of product efÞcacy, the 12AS formulation 12AS label rates, indicates a price ratio of WG:12AS at (Brown et al. 2001) was less efÞcient than the WG 1:1, if equivalent volumes of water are used to dilute formulation when applied against the container- the product. Given that problems with mixing large breeding species Ae. aegypti, Oc. notoscriptus (LC95 volumes of WG for ultralow volume applications have values converted to Bti ITU/liter, 421.07) and the been indicated previously (Muller et al. 2001), the saltmarsh species Oc. vigilax (LC95 values converted amount of water may need to be increased for proÞ- to Bti ITU/liter, 110.84). Such variation in product cient mixing, thus impacting on ferrying time and efÞcacy has been reported previously (Brown et al. overall efÞcacy of aerial treatments. However, the WG 2001, Fillinger et al. 2003) and may be related to formulation has shown advantages over the 12AS for- formulation characteristics. mulation not only because of its increased potency The test concentrations used in the small plot Þeld against a wide range of mosquito vectors but also trials were calculated after measuring the depth of the because the WG can be economically transported and water in the ponds to be 80 cm. This study demon- stored and has an extended shelf life. strates that presenting the results in this form provided the best indication of product concentration considering that product is dispersed throughout the water Acknowledgments column. In previous small-plot Þeld trials, Ragoonan- We thank Valent Biosciences for supplying product, and ansingh et al. (1992) and Fillinger et al. (2003), cal- Kay Marshall for producing and supplying larvae, Leaf Liai- culated product concentration assuming a standard sons for permission to conduct Þeld trial on property and water depth of 10 cm, but their ponds were actually Logan City Council for support. This work was funded by a deeper, thus overestimating product concentration. It Queensland Health Arbovirus Prevention Grant and the Lo- is difÞcult to directly compare our results with all cal Governments contributing to the Mosquito and Arbovirus other publications on VectoBac products, because ap- Research Committee, Inc. plication rates are expressed in many different ways (de Barjac 1990). The amount of product applied to mosquito habitats References Cited is usually calculated using surface area to be treated, Abbott. 1992. B. t. products manual. Abbott Laboratories, thus the concentration of product in ponds can vary Chicago, IL. depending on water depth. Our results indicate that Abbott, W. S. 1925. A method for computing the effective- the WG formulation loses little efÞcacy when diluted ness of an insecticide. J. Econ. Entomol. 18: 265Ð267. to concentrations equivalent to laboratory LC95 values Anscombe, F. J. 1948. The transformation of Poission, bio- in the Þeld. The registered label rate for WG is 125Ð500 nomial and negative bionomial data. Biometrika 35: 246Ð g/ha or 0.016Ð0.063 ppm in 80-cm deep water and for 254. 12AS, 0.3Ð1.2 liters/ha or 0.04Ð0.16 ppm in 80-cm deep Becker, N., M. Zgomba, M. Ludwig, D. Petric, and F. Rettich. 1992. Factors inßuencing the activity of Bacillus thurin- water. For treatment of mosquito habitats at depths giensis var. israelensis treatments. J. Am. Mosq. Control greater than this, the concentration of product applied Assoc. 8: 285Ð289. at registered label rates would be approaching those Boyd, A. M., and B. H. Kay. 1999. Experimental infection doses where suboptimal control was observed in the and transmission of Barmah Forest virus by Aedes vigilax laboratory. When selecting an appropriate application (Diptera: Culicidae). J. Med. Entomol. 36: 186Ð189. 1790 JOURNAL OF ECONOMIC ENTOMOLOGY Vol. 96, no. 6

Boyd, A. M., and B. H. Kay. 2000. Vector competence of Hershey, A. E., L. Shannon, R. Axler, C. Ernst, and P. Mick- Aedes aegypti, Culex sitiens, Culex annulirostris, and Culex elson. 1995. Effects of methoprene and Bti (Bacillus thu- quinquefasciatus (Diptera: Culicidae) for Barmah Forest ringiensis var. israelensis) on non-target insects. Hydro- virus. J. Med. Entomol. 37: 660Ð663. biologia 308: 219Ð227. Brown, M. D., D. Thomas, and B. H. Kay. 1998a. Acute Karch, S., Z. A. Manzambi, and J. J. Salaun. 1991. Field trials toxicity of selected pesticides to the PaciÞc blue-eye, with VectoLex (Bacillus sphaericus) and VectoBac (Ba- Pseudomugil signifer (Pisces). J. Am. Mosq. Control As- cillus thuringiensis (H-14)) against Anopheles gambiae soc. 14: 463Ð466. and Culex quinquefasciatus breeding in Zaire. J. Am. Brown, M. D., D. Thomas, K. Watson, and B. H. Kay. 1998b. Mosq. Control Assoc. 7: 176Ð179. Laboratory and Þeld evaluation of efÞcacy of VectoBac Kay, B. H. 1982. Three modes of transmission of Ross River 12AS against Culex sitiens (Diptera: Culicidae) larvae. virus by Aedes vigilax (Skuse). Aust. J. Exp. Biol. Med. Sci. J. Am. Mosq. Control Assoc. 14: 183Ð185. 60: 339Ð344. Brown, M. D., D. Thomas, P. Mason, J. G. Greenwood, and Kay, B. H., I. D. Fanning, and J. G. Carley. 1984. The vector B. H. Kay. 1999. Laboratory and Þeld evaluation of the competence of Australian Culex annulirostris with Mur- efÞcacy of four insecticides for Aedes vigilax (Diptera: ray Valley encephalitis and Kunjin viruses. Aust. J. Exp. Culicidae) and toxicity to the nontarget shrimp Leander Biol. Med. Sci. 62: 641Ð650. tenuicornis (Decapoda: Palaemonidae). J. Econ. Entomol. Kroeger, A., O. Horstick, C. Riedl, A. Kaiser, and N. Becker. 92: 1045Ð1051. 1995. The potential for malaria control with the biolog- Brown, M. D., T. M. Watson, S. Green, J. G. Greenwood, D. ical larvicide Bacillus thuringiensis israelensis (Bti)in Purdie, and B. H. Kay. 2000. Toxicity of insecticides for Peru and Ecuador. Acta Trop. 60: 47Ð57. control of freshwater Culex annulirostris (Diptera: Cu- McDonald, G., I. R. Smith, and G. P. Sheldon. 1977. Labo- licidae) to the nontarget shrimp, Caradina indistincta ratory rearing of Culex annulirostris Skuse (Diptera: Cu- (Decapoda: Atyidae). J. Econ. Entomol. 93: 667Ð672. licidae). J. Aust. Entomol. Soc. 16: 353Ð358. Brown, M. D., J. Carter, T. M. Watson, P. Thomas, G. San- Mulla, M. S., H. A. Darwazeh, and M. Zgomba. 1990. Effect taguliana, D. M. Purdie, and B. H. Kay. 2001. Evaluation of some environmental factors on the efÞcacy of Bacillus of liquid Bacillus thuringiensis var. israelensis products for sphaericus 2362 and Bacillus thuringiensis H-14 against control of Australian Aedes arbovirus vectors. J. Am. mosquitoes. Bull. Soc. Vector Ecol. 15: 166Ð175. Mosq. Control Assoc. 17: 8Ð12. Muller, M., S. Ritchie, P. Foley, and M. Brown. 2001. A new Brown, M. D., J. Carter, D. Thomas, D. M. Purdie, and B. H. Bti formulation for mosquito control: comparisons of Vec- Kay. 2002. Pulse-exposure effects of selected insecti- toBac 12AS liquid and VectoBac water dispersible gran- cides to juvenile Australian crimson-spotted rainbowÞsh ule. Arbovirus Res. Aust. 8: 256Ð262. (Melanotaenia duboulayi). J. Econ. Entomol. 95: 294Ð298. Piegorsch, W. W., and A. J. Bailer. 1997. Statistics for envi- Brown, M. D., T. M. Watson, J. Carter, D. Purdie, and B. H. ronmental biology and toxicology. Chapman & Hall, Lon- Kay. 2004. Toxicity of VectoLex (Bacillus sphaericus) don, England. products to selected Australian mosquito and non-target Pusztai, M., P. Fast, L. Gringorten, H. Kaplan, T. Lessard, and species. J. Econ. Entomol. (in press). P. R. Carey. 1991. The mechanism of sunlight-mediated de Barjac, H. 1990. ClassiÞcation of Bacillus sphaericus inactivation of Bacillus thuringiensis crystals. J. Biochem. strains and comparative toxicity to mosquito larvae, pp. 273: 43Ð48. 228Ð236. In H. de Barjac and D. J. Sutherland (eds.), Ragoonanansingh, R. N., K. J. Njunwa, C. F. Curtis, and N. Bacterial control of mosquitoes and black ßies. Rutgers Becker. 1992. A Þeld study of Bacillus sphaericus for the University Press, New Brunswick, NJ. control of culicine and anopheline mosquito larvae in Fillinger, U., B. G. Knols, and N. Becker. 2003. EfÞcacy and Tanzania. Bull. Soc. Vector Ecol. 17: 45Ð50. efÞciency of new Bacillus thuringiensis var. israelensis and Regis, L., M. H. Silva-Filha, C. Nielsen-LeRoux, and J. F. Bacillus sphaericus formulations against Afrotropical Charles. 2001. Bacteriological larvicides of dipteran dis- anophelines in Western Kenya. Trop. Med. Int. Health. 8: ease vectors. Trends Parasitol. 17: 377Ð380. 37Ð47. Russell, R. C. 1995. Arboviruses and their vectors in Aus- Finney, D. J. 1971. Probit analysis. Cambridge University tralia: an update on the ecology and epidemiology of Press, Cambridge, United Kingdom. some mosquito-borne arboviruses. Rev. Med. Vet. Ento- Gelernter, W., and G. E. Schwab. 1993. Transgenic bacteria, mol. 83: 142Ð158. viruses, algae and other microorganisms as Bacillus thu- Russell, R. C. 1998. Vectors vs. humans in Australia - Who is ringiensis toxin delivery systems, pp. 89Ð104. In P. En- on top down under? An update on vector-borne diseases twistle, M. Bailey, J. Cory, and S. Higgs (eds.), Bacillus and research on vectors in Australia J. Vector Ecol. 23: thuringiensis israelensis, an environmental bio-pesticide: 1Ð46. theory and practice. Wiley, Sussex, United Kingdom. Ryan, P. A., K. A. Do, and B. H. Kay. 2000. DeÞnition of Ross Georghiou, G. P., and M. C. Wirth. 1997. Inßuence of ex- River virus vectors at Maroochy Shire, Australia. J. Med. posure to single versus multiple toxins of Bacillus thurin- Entomol. 37: 146Ð152. giensis subsp. israelensis on development of resistance in SAS Institute. 2001. SAS online doc. computer program, the mosquito Culex quinquefasciatus (Diptera: Culici- version 8.2. SAS Institute, Cary, NC. dae). Appl. Environ. Microbiol. 63: 1095Ð1101. Watson, T. M., and B. H. Kay. 1998. Vector competence of Glare, T., and M. O’Callaghan. 2000. Assessing toxicity to Aedes notoscriptus (Diptera: Culicidae) for Ross River insects, pp. 27Ð29. In J. Wiley (ed.), Bacillus thuringiensis: virus in Queensland, Australia. J. Med. Entomol. 35: 104Ð biology, ecology and safety. Wiley, West Sussex, United 106. Kingdom. Watson, T. M., and B. H. Kay. 1999. Vector competence of Goldberg, L., and J. Margalit. 1977. A bacterial spore dem- Aedes notoscriptus (Diptera: Culicidae) for Barmah For- onstrating rapid larvicidal activity against Anopheles ser- est virus and of this species and Aedes aegypti (Diptera: gentii, Uranotaenia unguciulata, Culex univitattus, Aedes Culicidae) for dengue 1Ð4 viruses in Queensland, Aus- aegypti, and Culex pipiens. Mosq. News. 37: 355Ð358. tralia. J. Med. Entomol. 36: 508Ð514. December 2003 RUSSELL ET AL.: EFFICACY OF Bti FORMULATIONS AGAINST AUSTRALIAN MOSQUITOES 1791

Watson, T. M., K. L. Marshall, and B. H. Kay. 2000. Colo- Yousten, A. A., F. J. Genthner, and E. F. Benﬁeld. 1992. Fate nization and laboratory biology of Aedes notoscriptus from of Bacillus sphaericus and Bacillus thuringiensis serovar Brisbane, Australia. J. Am. Mosq. Control Assoc. 16: 138Ð israelensis in the aquatic environment. J. Am. Mosq. Con- 142. trol Assoc. 8: 143Ð148. [WHO] World Health Organization. 1981. Instruction for Zar, J. H. 1999. Multiple comparisons, pp. 208Ð214, In Bio- determining the susceptibility or resistance of mosquito statistical analysis. Prentice-Hall, Englewood Cliffs, NJ. larvae to insecticides. WHO/VBC/81.807, p. 6. World Health Organization, Geneva, Switzerland. Received for publication 16 June 2003; accepted 8 August 2003. ARBOVIRUS RESEARCH IN AUSTRALIA – VOLUME 9

THE USE OF BACILLUS THURINGIENSIS VAR. ISRAELENSIS FOR THE CONTROL OF ARBOVIRUS VECTORS IN THE AUSTRALIAN ENVIRONMENT.

Tanya Russell and Brian Kay

Queensland Institute of Medical Research, Australian Centre for International and Tropical Health and Nutrition and University of Queensland, Brisbane, Qld, Australia

INTRODUCTION

In Australia, the transmission of vector borne disease is of major concern. Ross River and Barmah Forest viruses, the primary arboviruses transmitted, can lead to the onset of debilitating endemic polyarthritis (Russell and Kay 2004). Annual activity of endemic polyarthritis has been recorded throughout Australia for over a century and has become increasingly frequent in recent times, affecting thousands of individuals during epidemic periods and causing significant impacts to the community in terms of morbidity, health costs and lost productivity (Kelly-Hope et al. 2004). In the absence of vaccines for such diseases, the community is reliant on mosquito control and personal protection to reduce human contact with mosquitoes and associated disease transmission. The primary focus of control operations in Australia is the point source reduction of larval populations. In contrast to worldwide procedures, a limited amount of adulticiding is conducted and only considered for epidemic responses (CLAG 2004, NEMMO 2004). The practice of larviciding is considered to be more economical and effective when compared with adulticiding, particularly when treating large areas, as larvae can be targeted while concentrated in pools before dispersing as adults (WHO 1997). There are three broad classes of larval habitats: freshwater ground-pools, container habitats and saline/brackish pools. As each habitat type is colonised by the larvae of specific mosquito species, mosquito control operations must be tailored to suit each habitat class. The aim of this paper is to review current control options for each habitat type, focusing on the application of Bacillus thuringiensis var. israelensis (Bti) for larval control. Furthermore, our recent trials assessing the use of Bti for the control of larvae in each habitat type are reviewed.

Bacillus thuringiensis var. israelensis Bti is an aerobic, gram-positive bacterial pathogen, initially described and identified as toxic to mosquito larvae by Goldberg and Margalit (1977). The activity of Bti is attributed to 4 crystallised proteins, which after ingestion, are dissolved and activated under the alkaline environment of the larval midgut and intestinal proteinases (Federici et al. 1990). This action releases toxins that bind to receptors on the epithelial midgut cells, resulting in an osmotic imbalance across the cell membranes, swelling and lysis of the affected cells, neutralising the alkalinity of the midgut which causes eventual insect death (Federici et al. 1990). Many factors influence the efficacy of microbial insecticide treatments. Of major importance is the feeding rate of larvae, since the activity of Bti is reliant on ingestion. Feeding rate can be influenced by 1) larval stage, 2) water temperature, 3) level of available food and 4) density of larvae. Firstly, the larvae are most susceptible as younger instars. When larvae mature to fourth instars, feeding activity declines and larvae are less likely to ingest a lethal dose in a short period of time. For this same reason, the non-feeding pupal stage is immune to Bti applications (Mulla 1990). Secondly, as water temperature decreases so does feeding activity and the consequential intake of microbial insecticides declines. Becker et al. (1992) found Aedes vexans (Meigen) larvae to be 10 times more sensitive to Bti at 25°C than at 5°C. Thirdly, as the level of available food increases the efficacy of Bti treatments decreases. Mulla et al. (1990) found that at the same concentration of 750 Bti ITU/L, 95% mortality of Culex quinquefasciatus Say was achieved in clear water and no mortality occurred in polluted water, because dense phytoplankton populations provided an alternative food source. Fourthly, as the density of larvae increases, competition for food increases and so Bti dosages should be increased (Becker et al. 1992). Strong sunlight appears to degrade toxins and reduce Bti activity. Becker et al. (1992) found that the concentration of Bti required for 90% mortality of Aedes aegypti (L.) at 48 hours was 3 times higher in sunlight compared to bioassays conducted in shade.

Freshwater ground-pool habitats Freshwater ground-pool habitats include permanent and temporary (rain filled) water bodies, e.g. lakes, swamps, streams, wheel ruts and dams. The major species found in these habitats are Culex annulirostris Skuse and Cx. quinquefasciatus (Lee et al. 1989). Large numbers of Ochlerotatus procax (Skuse), Ochlerotatus normanensis (Taylor), Ochlerotatus sagax (Skuse), Ochlerotatus bancroftianus (Edwards), Ochlerotatus vittiger

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(Skuse) and Coquillettidia linealis (Skuse) are also produced following rains (Lee et al. 1984). The larvae of Cx. annulirostris, will inhabit almost any form of naturally occurring freshwater ground-pools. This species inhabits permanent water during the dry season and rapidly exploits shallow vegetated pools within one day of formation after rainfall (McDonald and Buchanan 1981, Watson and Kay 1999, Dale and Morris 1996) as well as water associated with irrigation, ricefields or sewage effluent disposal (Marks 1982, Kay et al. 1992). Culex quinquefasciatus on the other hand, prefers to breed in polluted or highly organic habitats, as well as in artificial containers (described below). The larvae of this species colonise creeks and dams that permanently retain water and are polluted by sewage effluent (Marks 1982). In contrast, the larvae of Oc. procax prefer freshwater sedges and tea-tree swamps that form along the coastline after periods of rainfall (Ryan and Kay 2000). Due to emerging evidence that these species are prominent vectors of Australian arboviruses, there has been a renewed emphasis on control of freshwater species. However, except for Cx. annulirostris and Cx. quinquefasciatus, there is a paucity of data on effective control options. At present, control measures are attempted via ground-based larvicide treatments or habitat modification and local councils also rely on public education to minimise exposure to these mosquito species. Approximately 1,500 L of liquid Bti is used to control mosquito larvae in temporary ponds and rainwater drainage areas in south-east Queensland per season (CLAG 2004, NEMMO 2004). However, in areas where there is dense macrophyte growth on the marginal edges of freshwater habitats, adequate foliage penetration of liquid Bti is difficult to achieve and granular s-methoprene formulations, predominately Altosid pellets, are often applied. Currently in Australia, no granular Bti formulations are registered for use against mosquitoes in freshwater habitats. Davidson et al. (1981) found that a commercial powder formulation of Bti was effective against Cx. annulirostris and Cx. quinquefasciatus in the laboratory and field within 48 hours. Two other publications assessed the use of Bti against ground-pool freshwater larvae in Australia, both conducted in the laboratory against Cx. annulirostris. Brown et al. (2001) found VectoBac 12AS to be highly effective and similarly Muller et al. (2001) recorded the VectoBac WG formulation to be extremely efficient. With this latter product, the Bti toxins are formulated onto micropellets that disperse rapidly when placed in water. The product is designed to be mixed on-site and applied as a liquid formulation (Abbott 1992). In an effort to provide information regarding Bti applications in freshwater ground-pool habitats, we continued to assess the use of VectoBac WG in the field. A small-plot field trial was conducted in Logan Shire, southeast Queensland to assess the field efficacy of VectoBac WG against Cx. annulirostris (Russell et al. 2003). In independent pools, five replicates of 100 caged larvae were exposed to six concentrations of the WG formulation and three control replicates remained untreated (Fig 1). To test duration of control, weekly additions of 100 larvae were made to each treated and untreated control bioassay cage. As with the initial cohort, mortality was assessed at 48 h post introduction. VectoBac WG was effective at controlling Cx. annulirostris in the field with mortality of third-instars ranging from 85 ± 3.6% to 100.0 ± 0% in pools treated at rates between 31 and 1000 g/ha (0.004 and 0.13 ppm). No residual activity of the treatment was recorded at 1 week post treatment (mortality in treated ponds equalled 4.0 ± 6.9%). Our results indicate that VectoBac WG is highly effective for the short term control of larvae in freshwater ground-pool habitats. Furthermore, it has additional advantages over the traditional liquid formulations in that it can be economically transported and stored with an extended shelf life.

Figure 1. Third-instar Cx. annulirostris were exposed to VectoBac WG treatments in specially designed predator exclusion cages (left) and replicates were placed in a series of isolated pools (right).

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Container habitats The major species inhabiting natural and artificial containers holding freshwater are Ae. aegypti (Lee et al. 1987), Ochlerotatus notoscriptus (Skuse) (Lee et al. 1984) and Cx. quinquefasciatus (Lee et al. 1989). In Australia, most of our knowledge regarding container inhabiting mosquitoes is derived from studies of the dengue vector Ae. aegypti in northern Queensland. The larvae of this species have been recorded in artificial containers e.g. rainwater tanks, roof gutters, pot plant bases, tyres and rubbish (Tun-Lin et al. 1995, Montgomery and Ritchie 2002), natural containers e.g. fallen palm fronds and bromeliad leaf axils (Ritchie and Broadsmith 1997) as well as subterranean sites e.g. mine shafts, service pits and wells (Russell et al. 1997). A major operational concern is identifying containers that produce disproportionate numbers of larvae, termed key containers. In several north Queensland localities, Russell et al. (1997) identified the importance of subterranean sites as key containers, particularly during the winter months, when although they accounted for only 15% of the total number of positive containers, they produced 97% of the total standing crop of larvae. During the summer months, Montgomery and Ritchie (2002) identified roof gutters to be key larval containers in Cairns. During the study, roof gutters accounted for 3% of the total larval containers, yet produced 52% of the total standing crop of larvae. Similarly, Oc. notoscriptus immatures have been recorded in a diverse range of container habitats, including artificial (Marks 1982, Watson 1998), natural (Fanning et al. 1997, Ritchie and Broadsmith 1997) and subterranean (Russell et al. 1997, Watson 1998) containers. It has been noted that the species prefers organically rich water in semi-shaded or shaded positions (Watson 1998). In Brisbane, Watson (1998) found natural containers (bromeliad plant axils and tree-holes), garden accoutrements (plant pots and pot plant bases) and rubbish (disposable containers, drink cans etc) to account for 81% of larval productivity. During summer, the most productive containers were discarded household items (garbage bins, buckets, kitchen items etc) producing 32% of larvae, and during winter, discarded household items and garden accoutrements produced 51% of larvae. As with ground-pool habitats, the control of container breeding species has proved largely ineffective to date. This is due to the complexity of the mosquito species involved and their cryptic larval habitats, but also due to the lack of motivation by householders. House to house inspections and health education are the main methods used for control. Although, Queensland legislation provides heavy fines for those breeding mosquitoes, community participation is low and there has been a general reluctance by politicians to apply such legislation (Larson et al. 1997). In north Queensland, where surveillance is carried out by Queensland Health, containers that cannot be emptied are treated with s-methoprene, rather than Bti, which has a limited role in controlling container breeding species (Queensland Health 2000). At present, the only product available for application by domestic householders, is a granular s- methoprene formulation available at hardware stores. The mode of action of s-methoprene, an insect growth regulator, does not result in death of the larvae; rather it prevents pupal emergence. This means that to monitor application performance, emergence of pupae must be recorded, a tedious task for householders when compared to simply observing larval death. Thus, Bti is seen as a more desirable product to market for the promotion of community participation, however minimal research has been conducted in this field. Preliminary laboratory trials conducted in Australia by Canyon (1997) and Brown et al. (2001) demonstrated that liquid formulations of Bti were effective against Ae. aegypti and Oc. notoscriptus. However, this work has not been extended into the field. A handful of promising studies have been conducted overseas, where residual activity of Bti was recorded in containers in the field 1 week post treatment (see Becker et al. 1991, Batra et al. 2000, Seleena et al. 2001). We conducted a case study, focusing on control in pot plant saucers, to assess the potential of Bti for the control of Ae. aegypti and Oc. notoscriptus. Preliminary trials were conducted, with the aim of producing a product that could be applied to the soil surface by the householder, as a combined fertilizer/insecticide treatment, which would leach through the soil to kill larvae in the basal saucer. However, the results indicated limited control of Ae. aegypti larvae when exposed to leachate of VectoBac G in the laboratory probably because the active ingredient bound strongly to the soil particles. Various Bti formulations (VectoBac WG, 12AS and G) were also added directly to the pot plant saucers. In the laboratory, liquid VectoBac 12AS proved to be the most effective product. Effective control (>80% mortality of target larvae) of Ae. aegypti exposed to a high dosage of VectoBac 12AS (0.3 ml/saucer) was recorded for 11 weeks post treatment and for 8 weeks post treatment of Oc. notoscriptus exposed to a lower dosage of VectoBac 12AS (60 µl/saucer). However, when this trial was replicated in the field under UV radiation, the residual control of the products was significantly reduced. In the field, VectoBac G proved to be the most effective product, but effective control of Oc. notoscriptus was not recorded after 1 week post treatment. These results highlight the influence of UV degradation on the residual activity of Bti formulations, but could suggest potential for use in shaded situations.

Saline/brackish pools Saline/brackish pools are formed in coastal intertidal wetland communities. As a consequence of the changing tidal cycles, intertidal wetlands are waterlogged at times and exposed at others. The substratum of intertidal wetlands is relatively flat, with shallow depressions separated by low mounds, often vegetated. As the

339 ARBOVIRUS RESEARCH IN AUSTRALIA – VOLUME 9 water drains from the wetlands with the receding tide, water often remains in these depressions for a period of time, forming stagnant pools, particularly in the saltmarsh and upper mangrove (Adam 1994). These isolated pools are colonised by the larvae of a range of mosquito species, often at very high densities. The primary species found in Queensland, New South Wales, Western Australia and the Northern Territory is Ochlerotatus vigilax (Skuse) (Kay and Jorgensen 1986, Webb and Russell 1999) and throughout the remainder of the country Ochlerotatus camptorhynchus (Thomson) predominates (Lee et al. 1984). Other species that are commonly associated with intertidal wetlands are Culex sitiens Wiedemann (Lee et al. 1989) and Verrallina funerea (Theobald) (Lee et al. 1987). The mosquito larvae inhabiting saline intertidal wetlands are subject to well organised, broadscale control in Queensland, Western Australia and Northern Territory and to a lesser extent in the remaining States. The protocols for control are well developed and precursors for larval hatching are regionally defined, e.g. in southeast Queensland, rainfall > 100 mm over a 24 h period or tidal amplitude > 2.45 m is used to indicate larval hatching. During the warmer months of the year, when mosquito breeding is prolific, large areas of saltmarsh and the upper mangrove zone are treated via aerial or ground based operations with Bti. Liquid formulations are the most commonly used and upwards of 50,000 L of liquid Bti is applied per season to south-east Queensland intertidal wetlands, covering an area of approximately 40,000 ha (CLAG 2004, NEMMO 2004). In Australia, liquid Bti formulations seem effective against Oc. vigilax (Mottram et al. 1989, Brown et al. 2001, Muller et al. 2001, Webb and Russell 2001) and Cx. sitiens (Brown et al. 1998, Muller et al. 2001) in the laboratory and field, as well as for Oc. camptorhynchus (Brown et al. 2001) and Ve. funerea (Jeffery et al. 2005) in the laboratory. A granular formulation (G), where Bti is carried on corn grit, has proved useful for applications in densely vegetated areas, e.g. mangrove areas, where liquid formulations have poor penetration. However, this product has a relatively low Bti ITU/mg rating and large amounts must be carried to effectively treat extensive saltmarsh areas at the registered label rate (7 kg/ha), resulting in expensive treatments. There are no economical alternatives available to treat heavily vegetated areas. Granular s-methoprene (Altosid pellets), is often used to treat vegetated areas and although it is relatively expensive, applications do provide residual control. As large densities of larvae are commonly found in pools of stagnant saline water in heavily vegetated areas, there is pressure from local council operators for the development of an economical and effective product that can be used in such areas (CLAG 2004). One approach to overcome the ‘bulk density’ problem associated with this product is to lower the registered label rate, thus reducing the amount of product required for treatment. To investigate this possibility, we conducted a series of field trials at Redland Shire, southeast Queensland, to evaluate its efficacy against Oc. vigilax. Initially, a small plot field trial was conducted, where caged third-instars were exposed to five dosages of VectoBac G (0.5 – 7 kg/ha), replicated in isolated saltmarsh pools and mortality was assessed at 48 hours post treatment (Fig 2). The results indicated that at dosages below 4 kg/ha, the efficacy of treatments became unreliable. Using this information, a large scale field trial was conducted where the product was applied using rotary-wing aircraft, with the target application rate of 5 kg/ha. Once again, cohorts of 100 caged third-instars were exposed to the VectoBac G treatment in isolated pools. The treatment proved to be effective with 100% mortality of Oc. vigilax recorded at 48 hours post treatment. The distribution of product over the treatment area was assessed using a series of catch trays arranged in a grid-like fashion and an average dosage of 5.7 kg/ha was recorded. However, the 60% coefficient of variation for the application indicated considerable variability in actual dosage at individual sites. For this reason we plan to extend these trials.

Figure 2. Recording mortality of caged Oc. vigilax larvae exposed to VectoBac G treatment in isolated saltmarsh pools.

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CONCLUSION

Communities will continue to view mosquito control as being important to minimise the public health and economic impacts caused by mosquito borne disease, and also to create reasonable quality of life. Consequently, there is an ongoing need for research to define insecticide characteristics to ensure they are used appropriately and efficaciously with minimal environmental impacts. Our results indicate that the various formulations of Bti can be effectively adapted for use against freshwater ground-pool, container and saline/brackish larvae. The findings highlight the importance of tailoring research efforts to provide information that can be adapted for the control of larvae in each of the broad habitat classes. The water dispersible granule (VectoBac WG) was found to be effective at low dosages against freshwater ground-pool larvae, whilst the liquid formulation (VectoBac 12AS) was the most effective against container inhabiting larvae in areas of minimal UV radiation. In saline/brackish pools, preliminary trials indicated that the granular formulation (VectoBac G) has potential to be a useful product for application in heavily vegetated areas.

ACKNOWLEDGEMENTS

We thank Kay Marshall, Scott Lyons and the members of the Mosquito Control Laboratory, Queensland Institute of Medical Research for technical support and Valent Biosciences who supplied product used during the trials. Regarding the freshwater ground-pool field trial, we thank Michael Brown for initiating the project, Leaf Liaisons for permission to conduct trials on their property and Logan City Council for their support. This work was part funded by a Queensland Health Arbovirus Prevention Grant and the Local Governments contributing to the Mosquito and Arbovirus Research Committee, Inc. Regarding the saline/brackish helicopter trial, we thank Redland Shire Council for funding as well as Darren Alsemgeest, George Santagiuliana and Scott Dunsdon and the technical staff of the Pest Control Services for assistance with field work. Analysis of product distribution and the theoretical application rate was undertaken by Greg Dorr at the Centre for Pesticide Application and Safety, The University of Queensland.

REFERENCES

Abbott (1992) B.t. products manual. Chicago: Abbott Laboratories. Adam P (1994) Saltmarsh and mangrove. In: Australian vegetation (Groves RH, ed), pp 395-435. Cambridge: Cambridge University Press. Batra CP, Mittal PK and Adak T (2000) Control of Aedes aegypti breeding in desert coolers and tires by use of Bacillus thuringiensis var. Israelensis formulation. J Am Mosq Control Assoc 16: 321-323. Becker N, Djakarta S, Kaiser A, Zulhasril and Ludwig HW (1991) Efficacy of a new tablet formulation of an asporogenous strain of Bacillus thuringiensis israelensis against larvae of Aedes aegypti. Bull Soc Vector Ecol 16: 176-182. Becker N, Zgomba M, Ludwig M, Petric D and Rettich F (1992) Factors influencing the activity of Bacillus thuringiensis var. israelensis treatments. J Am Mosq Control Assoc 8: 285-289. Brown MD, Thomas D, Watson K and Kay BH (1998) Laboratory and field evaluation of efficacy of VectoBac 12AS against Culex sitiens (Diptera: Culicidae) larvae. J Am Mosq Control Assoc 14: 183-185. Brown MD, Carter J, Watson TM, Thomas P, Santaguliana G, Purdie DM and Kay BH (2001) Evaluation of liquid Bacillus thuringiensis var. israelensis products for control of Australian Aedes arbovirus vectors. J Am Mosq Control Assoc 17: 8-12. Canyon DV (1997) High variability in the effectiveness of aqueous Bacillus thuringiensis israelensis (Bti) formulations against Aedes aegypti mosquitoes. Bull Mosq Control Assoc Aust 9: 21-27. CLAG (2004) Contiguous Local Authorities Group, Regional Mosquito Control Committee, Quarterly Reports. Dale PER and Morris CD (1996) Culex annulirostris breeding in urban areas: using remote sensing and digital image analysis to develop a rapid predictor of potential breeding areas. J Am Mosq Control Assoc 12: 316-320. Davidson EW, Sweeney AW and Cooper R (1981) Comparative field trials of Bacillus sphaericus strain 1593 and B. thuringiensis var. israelensis commercial powder formulations. J Econ Entomol 74: 350-354. Fanning ID, Crisp G and Mottram P (1997) Bromeliads as breeding sites for mosquitoes in south-east Queensland. Arbovirus Res Aust 7: 82-83. Federici BA, Luthy P and Ibarra JE (1990) Parasporal body of Bacillus thuringiensis israelensis: structure, protein composition, and toxicity. In: Bacterial control of mosquitoes and blackflies (de Barjac H, Sutherland DJ, eds). New Brunswick: Rutgers University Press. Goldberg L and Margalit J (1977) A bacterial spore demonstrating rapid larvicidal activity against Anopheles sergentii, Uranotaenia unguciulata, Culex univitattus, Aedes aegypti and Culex pipiens. Mosq News 37: 355-358.

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Jeffery JAL, Kay BH and Ryan P (2005) Know thine enemy - Biology and control of brackish water vectors, particularly Verrallina funerea (Theobald), in Maroochy shire, Queensland. Arbovirus Res Aust 9. 153- 158. Kay BH and Jorgensen WK (1986) Eggs of Aedes vigilax and their distribution on plants and soil in south east Queensland (Australia) saltmarsh. J Aust Entomol Soc 25: 267-272. Kay BH, Piper RG, Falk PE, Battistutta D, Fanning ID and Lisle AT (1992) Mosquitoes from ricefields at Mareeba, North Queensland, Australia. Gen Appl Entomol 24: 19-32. Kelly-Hope LA, Purdie DM and Kay BH (2004) Ross River virus disease in Australia, 1886-1998, with analysis of risk factors associated with outbreaks. J Med Entomol 41: 133-150. Larson A, Bryan JH and Howard P (1997) Communities' roles in mosquito-borne disease control: lessons from two Queensland studies. Arbovirus Res Aust 7: 141-146. Lee DJ, Hicks MM, Griffiths M, Russell RC and Marks EN (1984) The Culicidae of the Australasian region. Volume 3. Canberra: Australian Government Publishing Service. Lee DJ, Hicks MM, Griffiths M, Debenham ML, Bryan JH, Russell RC, Geary M and Marks EN (1987) The Culicidae of the Australasian region. Volume 4. Canberra: Australian Government Publishing Service. Lee DJ, Hicks MM, Debenham ML, Griffiths M, Marks EN, Bryan JH and Russell RC (1989) The Culicidae of the Australasian region. Volume 7. Canberra: Australian Government Publishing Service. Marks EN (1982) An atlas of common Queensland mosquitoes. In, p 75. Brisbane: Queensland Institute of Medical Research. McDonald G and Buchanan GA (1981) The mosquito and predatory insect fauna inhabiting fresh-water ponds, with particular reference to Culex annulirostris Skuse (Diptera: Culicidae). Aust J Ecol 6: 21-27. Montgomery BL and Ritchie S (2002) Roof gutters: A key container for Aedes aegypti and Ochlerotatus notoscriptus (Dipters: Culicidae) in Australia. Am J Trop Med Hyg 67: 244-246. Mottram P, Madill B and Ebsworth P (1989) Efficacy of Bacillus thuringiensis var. israelensis for control of saltmarsh mosquitoes. Arbovirus Res Aust 5: 185-188. Mulla MS (1990) Activity, field efficacy, and use of Bacillus thuringiensis israelensis against mosquitoes. In: Bacterial control of mosquitoes and black flies (de Barjac H, Sutherland DJ, eds). New Brunswick: Rutgers University Press. Mulla MS, Darwazeh HA and Zgomba M (1990) Effect of some environmental factors on the efficacy of Bacillus-sphaericus 2362 and Bacillus-thuringiensis H-14 against mosquitoes. Bull Soc Vector Ecol 15: 166-175. Muller M, Ritchie S, Foley P and Brown M (2001) A new Bti formulation for mosquito control: Comparisons of VectoBac 12AS liquid and VectoBac water dispersible granule. Arbovirus Res Aust 8: 256-262. NEMMO (2004) North East Moreton Mosquito Organisation, Quarterly Reports. Queensland Health (2000) Dengue fever management plan for North Queensland 2000 - 2005. In, p 57. Cairns: Queensland Government. Ritchie SA and Broadsmith G (1997) Operational and scientific notes: Efficacy of Altosid pellets and granules against Aedes aegypti in ornamental bromeliads. J Am Mosq Control Assoc 13: 201-202. Russell BM, Foley P and Kay BH (1997) The importance of surface versus subterranean mosquito breeding during winter in north Queensland. Arbovirus Res Aust 7: 240-242. Russell R C and Kay BH (2004) Medical entomology: changes in the spectrum of mosquito-borne disease in Australia and other vector threats and risks, 1972–2004. Aust J Entomol 43: 271-282. Russell TL, Brown MD, Purdie D, Ryan PA and Kay BH (2003) Efficacy of VectoBac (Bacillus thuringiensis var. israelensis) formulations for mosquito control in Australia. J Econ Entomol 96: 1786-1791. Ryan PA and Kay BH (2000) Emergence trapping of mosquitoes (Diptera: Culicidae) in brackish forest habitats in Maroochy Shire, south-east Queensland, Australia, and a management option for Verrallina funerea (Theobald) and Aedes procax (Skuse). Aust J Entomol 39: 212-218. Seleena P, Lee HL and Chiang YF (2001) Thermal application of Bacillus thuringiensis serovar israelensis for dengue vector control. J Vector Ecol 26: 110-113. Tun-Lin W, Kay BH and Barnes A (1995) Understanding productivity, a key to Aedes aegypti surveillance. Am J Trop Med Hyg 53: 595-601. Watson TM (1998) Ecology and behaviour of Aedes notoscriptus (Skuse): Implications for arbovirus transmission in southeast Queensland. PhD Thesis, School of Population Health, University of Queensland. Watson TM and Kay BH (1999) Review of freshwater control programs, unpublished report, February 1999. In: Local Authorities' Research Committee, 1989-1999 annual report (Watson TM, ed), pp 70-103. Brisbane: Queensland Institute of Medical Research. Webb CE and Russell RC (2001) Do we spray? Did it work? Indices for control of Aedes vigilax larvae in Homebush Bay. Arbovirus Res Aust 8: 387-390.

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Appendix B

Toxicity of three insecticides to the commercial prawns Penaeus japonicus and Penaeus monodon

B.1 Introduction

During the term of my PhD, I was commissioned by the Rocky Point Prawn Farm, Gold Coast, to investigate the toxicity of the insecticides used for mosquito control to their stock. The tidal water from saltmarshes and mangroves is sourced for their aquaculture ponds. The operators of the farm were concerned that insecticide residue in the water may impact on the survival of their stock. In response, the acute toxicity of biologically-based insecticides, Bti and s-methoprene, and the organophosphate, chlorpyrifos, to the commonly farmed prawn species Penaeus japonicus and Penaeus monodon was investigated.

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B.2 Methods

Acute toxicity studies were conducted against post-larvae of P. japonicus and P. monodon. Specimens were obtained from stock at the Rocky Point Prawn Farm, Gold Coast. To minimize variability of response to test material, post-larvae of uniform age (3 – 4 weeks) and size (carapace length ranging between 10 – 15 mm) were selected.

B.2.1 Acute toxicity trials

Concentration-response data was developed for the biologically-based insecticides: Bti (VectoBac 12AS; Abbott Laboratories) and s-methoprene (Altosid Liquid Larvicide; Novartis Animal Health Australasia) as well as the organophosphate chlorpyrifos (Deter Insecticide; AI: 500g/L chlorpyrifos; Bayer Cropscience).

Post-larvae were exposed to serial dilutions of each pesticide in filtered and UV sterilized seawater (33mg/L salinity). To eliminate physiological stress associated with translocation, water was obtained from the Rocky Point hatchery where the larvae were reared. Five replicates, each of 20 specimens, were introduced into beakers containing 200 ml of test concentration. Five beakers holding 20 post-larvae in 200 ml of untreated seawater were used as controls. Test specimens were individually removed from holding trays and distributed randomly among the test beakers. The larvae were not fed during the testing period to minimize variability caused by nutritional and metabolic condition.

Initially, a number of range-finding tests with widely spread exposure concentrations were conducted. Based on these tests, a narrow range of concentrations that straddled the effective range was conducted to determine LC50 and LC95 values. The assays were conducted at 26oC under a light:dark cycle of 14:10 hours. The numbers surviving were counted at 48 h. Death or lack of reaction to gentle prodding with a glass pipette, was the measured deleterious response. Appendix B 247

B.2.2 Statistical methods

The data were analysed using probit analysis based on the Normal model for log-linear mortality (Finney 1971, PROC PROBIT, SAS Institute 2001). This method was more appropriate than standard probit analysis, as the data were not normally distributed. This method was adopted in favour of Abbott’s formula (Abbott 1925) because it does not modify the exposure variable, and thus has negligible impact on the curve.

A safety margin of insecticide application was calculated by dividing the LC95 value by the respective estimated field concentration (EFC) of the insecticide, for each species. The EFC of the insecticides was calculated using the highest registered label rate (VectoBac 12AS 1.2 L/ha and ALL 360 ml/ha) when applied to a saltmarsh pool with an average depth of 15 cm. The EFC of Deter Insecticide was taken as the highest level of chlorpyrifos recorded in Australian surface waters (National Registration Authority 2000).

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B.3 Results

Both Penaeus sp. exhibited relatively similar levels of susceptibility to the insecticides tested. The biologically-based insecticides, Bti and s-methoprene, both exhibited a low acute toxicity to the Penaeus species. The LC95 values recorded for Bti were 12,371 and 14,493 ppm VectoBac 12AS for P. japonicus and P. monodon, respectively and for s-methoprene were 105.36 and 92.73 ppm AI for P. japonicus and P. monodon, respectively (Table B.1).

Bti was the least toxic insecticide, as the LC95 value was at least 14 thousand times greater than the EFC (0.86 ppm VectoBac 12AS). The safety margin of s-methoprene applications was at least 77 times greater than the EFC (1.2 ppb AI).

In contrast, the organophosphate chlorpyrifos was highly toxic, with LC95 values of 0.159 and 0.365 ppb AI recorded for P. japonicus and P. monodon, respectively (Table B.1). As these values represent only 1/166 and 1/71, respectively, of the EFC for chlorpyrifos (26 ppb; National Registration Authority 2000), exposure of P. japonicus and P. monodon to chlorpyrifos under field conditions is likely to result in significant mortality.

Appendix B 249

a .17±0.20 16,852 (df) Slope±SE SM 2 x 27.44) 121(44) 1.96±0.23 77 0.662) 133(67) 0.92±0.09 0.014 ortality of commercial prawns after 48 h exposure. (5,772<85,230) 256(73) 0.97±0.21 14,318 (83.35<156.23) 261(52) 2.71±0.41 87 (95% CL) 9(0.105<0.421) 175(31) 2.92±0.66 0.006 95 LC (95% CL) 50 LC 816.54(449.85<1,239) 14,493(7,394<51,968) 609(80) 1 / estimated field concentration 95 (ppm VectoBac 12AS) 389.89 (143.65 < 600.74) 12,371 (ppm) -methoprene(ppmAI) 30.49(22.95<36.79) 105.36 -methoprene(ppmAI) 16.71(11.69<21.4) 92.73(73.88<1 Bti s Chlorpyriphos (ppb AI) 0.050 (0.034 < 0.069) 0.15 Bti s Chlorpyriphos(ppbAI) 0.011(0.006<0.016) 0.365(0.230< Table B.1: Concentration of pesticides causing 50% and 95% m Insecticide tested Penaeus japonicus Penaeus monodon Notes a. Safety Margin = LC 250

B.4. Discussion

The Penaeus sp. tested in this study exhibited a low toxicity to the biologically-based insecticides, Bti and s-methoprene. However, the organophosphate, chlorpyrifos, was highly toxic to both Penaeus species. The amount of Bti and s-methoprene required to reach the

LC95 concentrations was extremely high and reduction of the abiotic water qualities was observed. It is likely that mortality after exposure to such high concentrations of Bti and s- methoprene was due to dissolved oxygen depletion rather than direct toxicity, as suggested by Snarski (1990).

The larvae of penaeids hatch as free-swimming, oceanic plankton that progress through various morphological stages, of increasing complexity, termed nauplius, protozoea and mysis. The larvae then moult into post-larvae that migrate into estuarine and coastal nursery grounds (Gore 1985, Dall et al. 1990, Rothlisberg 1998). This is the youngest stage that would be exposed in the field, and in the commercial sense (prawns are transferred from the hatchery to ponds as post-larvae), and were used in this study. The results indicate that the use of Bti and s-methoprene for mosquito control is unlikely to impact on the commercial prawn industry. There is a possibility that observed prawn die-offs could be related to organophosphate, namely chlorpyrifos, run-off from surrounding sugar-cane farms (where it is commonly used to control cane beetles).

However, it is difficult to extrapolate these laboratory results into a field situation where target animals as well as the insecticide may be confounded by a range of environmental influences. This is particularly the case for s-methoprene, whose success as a mosquito control agent depends on the hormone mimic interfering with normal neuroendocrine functions, and which often takes up to one week for impacts to be noted on the target organisms. Considering this, s-methoprene has potential to affect other organisms that undergo similar moult cycles; such impacts were not measured in this study.