Australian Plague Locust Reponse - Assessment of Effects
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Centre for Aquatic Pollution Identification and Management Technical Report #9
Bryant Gagliardi, Sara Long, Gavin Rose, Lisa Golding, Jason Lieschke, Tara Daw Quadros, Leon Metzeling, Vincent Pettigrove.
Table of Contents
- Executive Summary
- Study Objectives
- Materials and Methods
- Recommendations for Future Locust Control Programs
- Table 1: Summary of locust insecticide applications in Victoria on public land 2010-2011
- Table 2: Synthetic APLR-approved pesticides (DPI) and their half-lives and toxicity to aquatic organisms
- Table 3: Threatened species, their status and locations within expected the locust risk zone
- Table 4: APLR-approved insecticides measured above limit of detection in sediments.
- Table 5: Insecticides measured above limit of detection in surface waters.
- Table 6: Mean conductivity (EC), pH, dissolved oxygen (DO) and temperature of water at sites where in situ cages of invertebrates were deployed or used in laboratory exposures either before and/or after pesticide spraying
- Table 7: Abundance of Murray hardyhead captured by each gear type at each location surveyed.
- Table 8: Summary results from four catchments sampled in the SRA program.
- Table 9: Determination of impacted sediment samples contaminated with APLR-approved pesticides
- Table 10: Determination of impacted biomonitoring sites contaminated with APLR-approved pesticides in surface water
- Figure 1: Locust activity in eastern Australia in March 2010
- Figure 2: Locust activity in south eastern Australia in December 2010
- Figure 3: Field sampling sites
- Figure 4: Decision-making process to determine whether or not sites were impacted by the APLR
- Figure 5: Mean (± SE) percent survival in C. tepperi larvae following five day exposure to sediment from sites where APLR approved pesticides were detected in the sediment.
- Figure 6: Mean (± SE) percent difference in surviving larvae length compared to laboratory controls in C. tepperi larvae exposed for 5 days to sediment from sites where APLR-approved pesticides were detected in the sediment.
- Figure 7: Mean (± SE) adult emergence (normalized to laboratory control emergence) in C. tepperi larvae exposed for up to 15 days in sediment collected from sites where APLR-approved pesticides were detected in the sediment.
- Figure 8: Proportion (± SE) of deformed antennae in Chironomus larvae collected from locust pesticide contaminated sites, relative to reference.
- Figure 9: Proportion (± SE) of deformed antennae in Polypedilum larvae collected from an APLR-approved pesticide contaminated site (662), relative to reference.
- Figure 10: Mean survival of amphipods caged for 7-d on pre- and post-spray occasions at three locations as well as a reference site under laboratory control.
- Figure 11: Mean survival of snails caged for 7-d on pre- and post-spray occasions at three locations as well as a reference site and laboratory control and also exposed to the same site waters and sediments under laboratory conditions on the post spray occasion.
- Appendix 1: Study sites, sampling regime and endpoints assessed
- Appendix 2: Sites for EPA targeted macroinvertebrate monitoring
- Appendix 3: Limits of reporting (LOR) forAPLR approved insecticides
- Appendix 4: Organic carbon and organic matter levels for sediments contaminated with locust-registered pesticides
- Appendix 5: Total organic carbon (TOC) and Total Nitrogen levels for surface waters contaminated with locustregistered pesticides
- Appendix 6: Macroinvertebrate results from the EPA "targeted" sites sampled.
In spring 2010, Victoria was threatened with a potentially serious locust plague. In response, the Victorian Government launched the Australian Plague Locust Response (APLR) to mitigate locust impacts by biological and chemical control, headed by the Victorian Department of Primary Industries (DPI). The present study is the Ecotoxicological Monitoring component of the Audit, Produce and Environmental Monitoring (APEM) Project, which monitored the APLR for impacts on compliance with legislation, and for unwanted insecticide impacts.
The aim of this study was to monitor Victorian waterways in the plague locust risk zone for environmental impacts resulting from application of pesticides approved in the APLR. These results would in part inform best practices for future plague locust control with respect to pesticide applications. A total of 81 sites were monitored in this study. Presence of APLRapproved pesticides in environmental sediments and surface waters was determined by chemical analysis. Ecosystem impacts for each contaminated sample were inferred by assessing at least one of the following: toxicity of sediment to the freshwater midge Chironomus tepperi, inducement of morphological deformities in the freshwater midge Chironomidae by sediment and surface waters, and impacts to populations of the native endangered fish, the Murray hardyhead (Craterocephalus fluviatilis). Four river basins in the affected region were also monitored for impacts to macroinvertebrate communities, however this component of the study was compromised by flooding. Novel in situ cage bioassays were also trialled for three of the sites. These assessed the acute toxicity of sediments and surface waters to the freshwater snail Potamopyrgus antipodarum and the freshwater amphipod Austrochiltonia subtenuis.
Chemical analysis found 11 incidences of contamination by APLR-approved pesticides. Each incidence was found to either be occurring at a concentration not toxic to test organisms, or to be the result of the control of pests other than locusts. Hence, no evidence was found to suggest the APLR had caused aquatic ecosystem impairment in Victorian waterways.
The interpretation of this study with regard to recommendations for future locust plague control programs was complicated by atypical climatic events (including flooding and cool temperatures). These events probably lead to lower than forecast locust numbers, and fewer pesticide applications were conducted than predicted. This may be a contributing factor to the lack of aquatic impacts observed. However, the low number of APLR-approved pesticide detections in samples suggests that existing strategies to minimise entry of locust control pesticides into freshwater environments were successful. For this reason the authors suggest these measures be implicated in future locust control programs. Importantly, we also suggest better and timelier availability of pesticide use data, both on private and public land, to better and more promptly enable judgement as to whether or not detected pesticides are the result of locust control. Data should also be more specific in terms of volumes and types of chemicals used, as well as, timing and locations of applications. Finally, we suggest the inclusion of a thorough biomonitoring program, as in the present study, for future locust control programs.
Victorian locust plague
In spring 2010, Victoria experienced high densities of the pest species Australian plague locust (APL) Chortoicetes terminifera. This posed the threat of the worst locust plague in Victoria in 75 years; and had the potential to cause wide spread devastation to the Victorian agricultural sector.
Chortoicetes terminifera is a serious agricultural pest in Australia. An individual locust can consume 30 to 50 percent of its body weight daily, and swarms are very dense and highly mobile, migrating up to 700 km in a day. Swarms can be in excess of 50 individuals per square metre, consuming up to 10 tonnes of vegetation per day (Spratt, 2004). Typically pastures are at risk from APL damage, but crops (particularly winter cereals) may also suffer damage (Love and Riwoe, 2005).
Locusts (Orthoptera: Acrididae) are grasshoppers that demonstrate marked phenotypic changes in response to increased population density. At low densities, locusts are "solitarious" and avoid one another. However, upon crowding, locusts enter an aggregating, migratory "gregarious" phase. It is in this phase that locusts are potentially devastating to agriculture: both as massive bands of wingless juveniles ("nymphs" or "hoppers") migrating across land and as flying swarms of winged adults (Simpson and Sword, 2008).
The Australian Department of Agriculture, Fisheries and Forestry (DAFF) Australian Plague Locust Commission (APLC) reported high densities of locusts developing in New South Wales, including the Riverina region in March 2010 (Figure 1). These populations were largely the result of migrations from southwest Queensland and were sustained by favourable weather and rainfall conditions (DAFF, April 2010). By April 2010 a "major nymphal infestation" had been reported, and several southward migrations of adults resulted in an increase in reported locust densities in northwest, north central and northeast Victoria. Within Victoria, localised breeding and high-density egg-laying were observed, and serious nymphal infestations in October were consequently predicted. A high survival rate in these spring-emergent nymphs could potentially lead to a high-density locust "plague" of swarming adult locusts in these regions (DAFF, May 2010).
Hatching and fledging of locust nymphs; and subsequent swarming, occurred over spring 2010 and summer 2010-2011. This occurred at different times and different rates, depending on local soil types and climatic conditions of the various affected regions. Swarms across northern Victoria intensified, with swarms reported in the Bordertown-Ouyen, Cullulleraine-Mildura, Sea Lake-Swan Hill and Kerang-Echuca areas (see Figure 2).
There were several further significant and migratory events in November, including an apparent migration of locusts from northwest Victoria to the Hamilton region in southwest Victoria (DAFF, January 2011). This migration was unexpected and not forecast in the initial stages of the APLR.
Egg-laying was observed in the spring-emerging locusts, resulting in the emergence of a second generation in January. Swarms of second generation locusts were subsequently reported in the northwest, north central and northeast regions, and also now in the southwest region including the southern Grampians-Ararat area. High-density egg laying was predicted in this second generation of locusts, and eggs were predicted to go into diapause, potentially hatching in spring 2011 (DAFF, April 2010).
Cooler than average conditions slowed the hatching and development of locusts over the course of this event. Overcrowding of locusts in dense vegetation, preventing them from regulating their body temperature by basking in the sun, may also have lead to reduced populations (Millist and Abdalla, 2011). The Victorian locust plague was therefore much smaller than forecast, substantially reducing the volume of pesticides applied from that initially predicted.
Australian Plague Locust Response (APLR)
The DPI, through its Biosecurity Victoria Division (BV) commenced planning in autumn 2010 for the Australian Plague Locust Response (APLR). This response comprised two major components:
- Community and landholder engagement in control strategies, including government funding for landholder insecticide applications, and
- DPI BV coordinated official Emergency Response, including deployment of significant numbers of DPI staff through phased responses to the plague at regional control centres in Mildura, Horsham and Tatura.
DPI oversaw pesticide applications on public land, and private land managers were responsible for locust control on private land. The APEM outlined measures to minimise the risk of aquatic environmental impacts (DPI, 2010). For example, for public land spraying, the application of synthetic chemicals was forbidden within 1500 metres of waterways, or where water was expected within 72 hours (3 km in the case of RAMSAR wetlands). Where available, the biological control agent Metarhizium ("Green Guard") was to be used on public land, particularly in areas of "High Biodiversity Value". Metarhizium (the fungus Metarhizium anisopliae var. acridum) is an effective, specific treatment (Hunter et al., 2001) of Australian locusts which poses a very low risk to aquatic organisms (Milner et al., 2002). Treatment of locusts was also recommended at the nymphal stage, rather than the flying adult stage, to reduce volumes of pesticide used. Treatment of nymphs is more effective and efficient as locusts are less mobile at this stage; nymphs are concentrated in a small area, and, unlike adults, cannot easily avoid treatment by flying away (DPI, 2011a). Pesticides approved for usein the APLR are listed in Table 2 . Landholders who used these pesticides and/or Metarhizium for the control of locusts were eligible for government rebates under the Locust Insecticide Rebate Scheme (LIRS).
Summary pesticide application data was made available to the authors subsequent to field surveys and experiments. Public land spraying is summarised in Table 1. Approximately 888,426 ha of private land was sprayed with chemicals claimed on the LIRS (Alan Roberts, DPI, personal communication), and it is not known what area of land was treated by private landowners with chemicals not claimed on the scheme, or by pesticides not falling under the scheme.
|Chemicals Sprayed||# Cases||
|Green Guard SC1||119||1,617|
|Green Guard ULV2||5||3,370|
Monitoring aquatic environments for APLR-approved pesticide contamination and biological impacts
All synthetic pesticides approved for use in the APLR (hereby referred to as APLR-approved pesticides) are variously toxic to non-target invertebrates, as well as aquatic organisms (see Table 2). Agricultural pesticides may enter aquatic environments via spray drift, runoff or groundwater (Kookana et al., 1998), and have been previously detected in freshwater environments in Victoria (Rose et al., 2009). The effects of aquatic contamination (contaminant being defined as "a substance released by man's activities" (Moriarty, 1983)) on the ecosystem can be evaluated by ecotoxicology. Ecotoxicology is the study of contaminants and their effects on organisms in the environment (Newman, 1998).
Chemical runoff from anthropogenic activities can pose a threat to aquatic ecosystems, either through direct toxicity to sensitive organisms; or through indirect effects by alteration of abundances of pollution-sensitive and pollution-tolerant species (Marshall et al., 2010, Fleeger et al., 2003).
In any pesticide monitoring program, sediments are of particular interest as they often act as sinks for pollutants (Li et al., 2000). Sediment-bound contaminants can be resuspended by environmental events such as storms (Pitt, 1995), and can facilitate the entry of some chemicals into the food chain. Benthic macroinvetebrates, being in direct contact with sediments, are particularly at risk from sediment pollution. They can also act as an entry point for sediment-bound pollutants into the food chain (Lee et al., 2000).
|Class of pesticide||Name||Water solubility mg/L||Half life||Toxicity to aquatic organisms (water concentrations)|
|Organophosphate||Fenitrothion||30 (at 21 °C)||85 days (in sandy loam, under irradiation) (Dykes and Carpenter, 1988)||LC50 (96 h) bluegill sunfish 3.8mg/L, brook trout 1.7mg/L LC50 (48 h) carp 4.1 mg/L|
|Organophosphate||Chlorpyrifos||2 (at 25 °C)||60-120 days (in soil)||LC50 (96 hr) rainbow trout 0.003mg/L (24 hr) goldfish 0.18mg/L Toxic to crustaceans|
|Organophosphate||Diazinon||40 (at 20 °C)||2-4 weeks (in soil) (Wauchope et al., 1992)||LC50(96 hr) bluegill sunfish 16 mg/L, rainbow trout 2.6- 3.2 mg/L|
|Organophosphate||Maldison (Malathion)||145 (at 25 °C)||1-2 days (in aerobic soil)||LC50 rainbow trout 0.1 mg/L (MSDS: Maldison)|
|Carbamate||Carbaryl||40 (at 30 °C)||7-14 days (in sandy loam) 14-28 days (in clay loam)||LC50 (96 hr) rainbow trout 1.3 mg/L, bluegill sunfish 10 mg/L|
|Phenylpyrazole||Fipronil||2.4 (at pH 5) 2.2 (at pH 9) (Tomlin, 1997)||630 - 693 days (in aerobic soils) (Tomlin, 1997)||LC50 (96 h) rainbow trout 0.248 mg/L, European carp 0.430 mg/L, bluegill sunfish 0.085 mg/L, LC50 (48 h) daphnia 0.19 mg/L (MSDS: Fipronil)|
|Synthetic pyrethroid||Cyhalothrin||0.005 (at pH 6.5, 20 °C)||4-12 weeks (in soil)||LC50 bluegill sunfish 0.24 mg/L|
|Synthetic pyrethroid||Cyfluthrin||0.002 (at 20 °C)||2-16 days (soil photolysis, 28°C at pH 6.6; sandy loam) (Casjens, 2001)||LC50 (96h) golden orte 0.0032mg/L, rainbow trout 0.006mg/L, carp 0.022mg/L, bluegill sunfish 0.0015 mg/L|
|Synthetic pyrethroid||Cypermethrin||0.01 (at 25 °C)||ca. 13 weeks (in loamy soil)||LC50 (95h) rainbow trout 0.0028mg/L|
Importantly, aquatic ecotoxicology can verify or quantify the impacts of contaminated sediments and surface waters on biota. This approach gives ecological relevance to chemistry results. Ecotoxicology can ultimately indicate whether detected contaminants are having a deleterious effect on biota (at which point they are deemed "pollutants" (Moriarty, 1983)), or whether they are present at non-toxic concentrations and therefore not of concern. This is achieved through "biomonitoring", which is "the use of organisms to monitor contamination and imply possible effects on biota..." (Newman, 1998)
This study was conducted by the Victorian Centre for Aquatic Pollution Identification and Management (CAPIM), the Arthur Rylah Institute (ARI, the biodiversity research base for the Department of Sustainability and Environment [DSE]) and the Environmental Protection Authority, Victoria (EPA).
CAPIM conducted field surveys collecting sediment, water and chironomid larval samples for analysis; and conducted ecotoxicological bioassays on sediments and surface waters. ARI monitored several wetland populations of the threatened native fish species Murray hardyhead (Craterocephalus fluviatilis), which were known to occur within the locust risk zone. CAPIM and EPA Victoria shared responsibility for monitoring the health of several aquatic macroinvertebrate communities in the locust risk zone.
Field and laboratory toxicological assessments using test organisms: chironomids
Laboratory and field ecotoxicological analyses were carried out using the macroinvertebrate family Chironomidae (Diptera) ("non-biting midges") as test organisms. Chironomids are useful indicators of waterway pollution, particularly for sediments. Animals reside in the sediment where they settle as larvae after hatching and remain until they emerge as adults, spending a significant proportion of their lifecycle exposed to sediment-bound pollutants. The chironomid Chironomus tepperi has been used as a laboratory test species in toxicity testing in Australia (Choung et al., 2010, Kellar et al., 2011, Stevens et al., 2005). Laboratory tests measured a variety of endpoints, including lethal, sub-lethal acute and sub-lethal chronic.
Field and laboratory toxicological assessments using test organisms: trialling of snail and amphipod bioassays
Novel in situ and laboratory ecotoxicological experiments were also trialled at three of the 81 study sites. These used the test organisms Potamopyrgus antipodarum (an exotic freshwater snail) and Austrochiltonia subtenuis (a native freshwater amphipod). In situ bioassays involve exposing caged aquatic organisms to in-stream conditions, and form a vital link between laboratory based bioassays and field studies (Crane et al., 2007). One of the great advantages of in situ bioassays is the measurement of a time-integrated biological response that incorporates impacts from pulse events such as pesticide applications. These types of events are often difficult to capture in traditional laboratory-based bioassays, as grab collected samples can frequently miss episodic pulses of pollution. Freshwater snails and amphipods have been routinely used as in situ cage test organisms because of their robustness to cage conditions, ease of collection or culture, routine use in standardised laboratory protocols and wide range of sensitivity to toxicants (Schmitt et al., 2010, Schulz, 2003). There is a paucity of sediment toxicity data for most of the APLR-approved pesticides with respect to snails and amphipods. However species sensitivity differences do occur between commonly used agricultural pesticides (Ding et al., 2011), emphasizing the importance of using a range of test species to provide environmental protection. Together with field biomonitoring, laboratory bioassays and chemicals analyses, in situ cage bioassays provide a bridging piece of evidence to add to the overall weight of evidence approach in assessing effects of contaminants.
Monitoring for native fish impacts
Due to the potential for APLR-approved pesticides to impact native fish species, the Arthur Rylah Institute was contracted to monitor populations of threatened native fish in the locust risk zone. As well monitoring for APLR effects to threatened fish, this component of the study also provided a biomonitoring endpoint to assess APLR-induced toxicity to biota at monitored sites.
Several of the synthetic APLR-approved pesticides were considered to pose a potential risk to native fish stocks. Metarhizium has been classified as posing a low risk to fish (PRG, 2004), though this classification comes with a caveat stating that it is based on laboratory data or small field trials with indigenous species from areas outside of Australia. The US EPA has noted that Metarhizium can affect embryo and larval development within the Silversides family (Genthner and Middaugh, 1995), however, the environmental relevance of these results has been questioned (Milner et al., 2002). Two threatened fish species occurring in the locust risk zone, Murray hardyhead and unspecked hardyhead (Craterocephalus stercusmuscarum fulvus), are both members of the Silversides family whose breeding cycles would occur during the APLR. The toxicity of Metarhizium to any fish species located in the locust risk zone is not known, though it has been shown to have no adverse effects on the native eastern rainbow fish (Melanotaenia duboulayi) (Milner et al., 2002), a species of the same genus as the crimson-spotted rainbowfish (Melanotaemia fluviatilis), which is found within the locust risk zone.
Monitoring for macroinvertebrate community impacts
EPA Victoria was charged with monitoring several communities of aquatic macroinvertebrates within the locust risk zone. Macroinvertebrates (such as insects, snails and worms) are commonly used for monitoring of aquatic ecosystem health, for several reasons (EPA, 2003):
- They are ubiquitous: macroinvertebrates can be obtained from a wide range of aquatic systems and habitat types.
- They are relatively sedentary: macroinvertebrates, compared with other larger more mobile organisms such as fish; are relatively sedentary in nature, which ensures they are less likely to escape impacts.
- They have varying life histories ranging from weeks to years. This provides a cumulative indication of environmental conditions over time. It also allows for consideration of disturbances that take place at many different time scales. This is in contrast to groups that have a smaller range of life histories such as algae.
- The collection, identification and analysis of macroinvertebrates is relatively cost effective.
- Macroinvertebrates are the biotic indicator used in the Australian Rivers Assessment System (AusRivAS), a nation-wide program for monitoring stream health (Davies, 2000). This has resulted in a large body of knowledge on the taxonomy, ecology, and ecotoxicology of macroinvertebrates.
As most of the macroinvertebrates in streams are insects, there was also the potential for a response to the insecticides used in the APLR.
The aims of this study were:
- To determine the incidence of APLR-approved pesticides in monitored streams and wetlands in the Australian plague locust risk zone within Victoria
- To determine whether any such contamination was causing aquatic ecosystem impairment
- Where impairment was detected, to determine whether the contamination was a result of the APLR
- To trial novel laboratory and in situ snail and amphipod ecotoxicology bioassays
- To recommend, based on these findings, best practices for future locust control and associated environmental monitoring programs
Materials and Methods
Study area and sampling regime
Collection of samples for pesticide analyses and ecotoxicological bioassays
Three rounds of field sampling were undertaken (see Figure 3). Initially, selected sites (located within the locust risk zone in northern and north western Victoria) were to be sampled three times: prior to the commencement of public land spraying ("pre-spray", in order to provide reference samples for comparison, 28/9/10 – 27/10/10); after the approximate conclusion of public land spraying ("post-spray", 21/12/10 – 8/3/11); and the third round 2-3 months after the conclusion of spraying in order to monitor the ongoing health of sites.
Prior to the commencement of the third round of sampling, however, a change in study design was made. This was due to both the low detection rate of APLR-approved pesticides in the post-spray round of sampling, and the southerly migration of locusts. Instead of repeat monitoring the initial northern sites, this sampling round (19/4/11 - 15/6/11) mostly targeted south western Victoria. Sites found to have APLR-approved pesticides in the post-spray sampling round, however, were again sampled in this third round. Sites in the Wimmera region were re-sampled in the third round of sampling, as the results of pesticide analyses were not yet available at this time. 20 sites were sampled only in the second round of field sampling. These were the sites monitored by the EPA macroinvertebrate "targeted" monitoring (see Materials and Methods: Macroinvertebrate monitoring), and were sampled to provide chemical and ecotoxicological data to supplement this program. Three of the sites monitored by ARI for native fish populations were also sampled in order to supplement this program with chemical and ecotoxicological data. Spray periods were determined by regular communication with DPI. See Appendix 1 for all site names. Sites were selected to represent a range of land use types within the regions forecast to experience locust activity. A combination of riverine and wetland environments were selected that were thought likely to receive runoff from surrounding land. Site selection was informed in part by personal communication with several local expert bodies (North Central Catchment Management Authority [CMA], Wimmera CMA, Mallee CMA, Goulburn Broken CMA, Victorian DPI, Victorian DSE).
Site selection for snail and amphipod in situ and laboratory bioassay trials
Snail and amphipod bioassay trials were conducted at three of the 81 field sites. Cage deployment sites were selected to represent separate catchments at similar latitude where pesticide spraying was likely to occur as part of the APLR. The Victorian EPA database was used to confirm the prior occurrence of the biomonitoring species in the catchments to avoid accidental translocation of the invertebrates between catchments. Cages were deployed at a reference site and in the laboratory under controlled conditions were included to account for site habitat and water physico-chemical differences. The reference site was changed to another location after the pre-spray deployment due to low dissolved oxygen measured when retrieving the cages.
Threatened fish monitoring
From the initial proposed area to be sprayed, a list of populations of threatened fish to be monitored was determined (see Table 3).
|Common name||Scientific name||Location (catchments unless otherwise specified)||DSE Advisory threat status||FFG listed||EPBC|
|Silver perch||Bidyanus bidyanus||All catchments north of the Great Dividing Range||Critically endangered|
|Murray hardyhead||Craterocephalus fluviatilis||Kerang Lakes, Swan Hill and Mildura areas||Critically endangered||Listed|
|Unspecked hardyhead||Craterocephalus stercusmuscarum fulvus||All catchments north of the Great Dividing Range||Data Deficient||Listed||Vulnerable|
|Trout Cod||Maccullochella macquariensis||Broken and Goulburn||Critically endangered||Listed||Endangered|
|Murray cod||Maccullochella peelii peelii||All catchments north of the Great Dividing Range||Endangered||Listed||Vulnerable|
|Golden perch||Macquaria ambigua||All catchments north of the Great Dividing Range||Vulnerable|
|Macquarie perch||Macquaria australasica||Broken, Goulburn and Campaspe||Endangered||Listed||Endangered|
|Catfish||Tandanus tandanus||Loddon, Little Murray and Wimmera||Endangered||Listed|
Due to unseasonal weather patterns and extensive flooding, only the threatened Murray hardyhead was monitored. These unprecedented weather conditions also caused the Murray hardyhead sampling to be delayed from early December, to January, then again from January until February.
Four sites containing Murray hardyhead populations were sampled. These were: Site 613 (Woorinen North Lake) from the Swan Hill lakes, Site 614 (Round Lake, Lake Boga) from the Kerang lakes, and Site 612 (Cardross Lakes Basin 1) and Site 685 (Lake Koorlong) from the Mildura region. These sites were chosen as they represent the last remaining Murray hardyhead populations in Victoria, and were all in areas where the APLR was to occur.
EPA designed a two part monitoring program to detect if locust pesticides had an impact on aquatic macroinvertebrate communities.
The first part of the program aimed to gain added value from the pre-existing sampling program, the Sustainable Rivers Audit (SRA), which is co-funded by the Murray Darling Basin Authority and DSE. Four of the five river basins (Avoca, Broken, Campaspe and Wimmera) scheduled for sampling under this program during 2010/2011 were identified as being in the locust risk zone. These basins (up to 35 sites in each) were scheduled for sampling between October – December 2010 and as such could act as "pre-spray" samples. It was intended that information could then be obtained from DPI regarding which areas had been sprayed and hence which of these stream sites were likely to have been affected, and then carry-out repeat sampling which would then be carried out at 20 sites across these basins. This sampling would have indicated if there were any short-term, acute impacts. The follow-up autumn samples would then indicate if there were longer term affects of spraying on macroinvertebrate communities.
The second part of the program intended to take a more targeted approach, one that did not rely on SRA sampling but focussed on sampling streams in areas that had a high likelihood of being sprayed. These sites were determined by using maps of APL distribution. A total of 20 APL monitoring sites (10 site pairs all within 1km of each other) (see Figure 3 for site locations, Appendix 2 for site names) were selected to be sampled before and after spraying. The targeted streams were sampled during 18-20 October 2010 in the Avoca, Loddon and Wimmera basins, prior to any spraying in the area. In addition to the follow-up macroinvertebrate sampling, sediment and water samples were to be collected by CAPIM for chemical analyses.
Collection of sediment and surface samples.
Surface sediments (approximately top 2 cm) were collected with either a dip net or shovel, and wet filtered through a 63μm nylon sieve on site. This filtration was conducted in accordance with Australian and New Zealand water quality guidelines to remove the possible diluent effect of coarser particles (ANZECC/AMRCANZ, 2000). Fine sediments are also of particular relevance as particles less than 63μm are common in the gut of sediment-ingesting biota (Tessier et al., 1984). Filtered material was collected in a rinsed polyvinyl chloride bucket, returned to the laboratory and stored at approximately 4°C overnight to settle. Supernatant water was decanted, and settled sediment was stored in solvent-rinsed jars at 4°C until analyses and bioassays. One litre of surface water was collected on site in solvent-rinsed amber glass bottles, and stored at approximately 4°C until analyses.
Collection of chironomid larvae for deformity analysis
Larvae were collected on-site either with a 250 μm macroinvertebrate dip net, or by collecting surface sediments with a shovel, filtering this material with 500 μm nylon net and collecting remnant coarse material. Material from either the 250 μm or 500 μm net was transferred so a sorting tray, and Chironomus and Polypedilum larvae were live picked using forceps. They were transferred into 70-100% ethanol, and returned to the laboratory where they were stored at 4°C until analysis.
Collection of threatened fish
Fifty Murray hardyhead were collected with a seine net from Site 612 (Cardross Lakes Basin 1) on 12th October 2010 with an additional 100 Murray hardyhead collected from Site 614 (Round Lake, Lake Boga) on 19 and 20 January 2011. All fish were transferred to the aquaculture facilities at the Murray-Darling Freshwater Research Centre (MDFRC) in Mildura for population security and for breeding purposes.
Fish surveys were undertaken during the week of 21st February 2011 using both active (electrofishing and seine netting) and passive (fyke netting) techniques. Site conditions in some cases precluded the use of one (seine netting) or both active techniques (seine netting and electrofishing).
Boat electrofishing was conducted at Site 685 (Lake Koorlong) (1800 electrofishing seconds on-time) and Site 613 (3000 electrofishing seconds on-time) using a prototype EL65IIGEH with SK1 Switchbox (Grassl) generator with 13 kW output power. Seine netting was conducted at Site 613 using large and small mesh size nets with two sampling passes. Fyke netting was conducted at all four sites using four double winged fyke nets which were set late in the afternoon and retrieved the following morning. Soak times for the fyke nets varied between 16 ½ hours and 18 hours.
Standard EPA sampling procedures were employed (EPA, 2003) with two edge habitats sampled at all sites in both the SRA program and the targeted sites approach. Samples were collected using a sweep net from littoral areas of the stream, in beds of macrophytes, around logs and in backwaters.
Subsequent to the pre-spray sampling of targeted and SRA sites, field sampling was abandoned. This was primarily because flooding that had occurred in Victoria had meant any detected impact to macroinvertebrate communities could be attributed to either locust spraying or the massive disturbance caused by the flooding. Secondly, as there was no information available about where and when spraying had occurred on private land, selection of sites from the SRA program for follow-up sampling was problematic with little confidence that the sites chosen would be representative of pesticide impacts from private spraying.
The program was amended so that any follow-up macroinvertebrate sampling would occur if pesticide of concern were found at any of the targeted sites. This would be based on sediment sampling conducted by CAPIM. The result of CAPIM sediment and water sampling and analysis found only a single detection of an APLR-approved pesticide. On this basis, EPA decided to not conduct any follow-up macroinvertebrate sampling at any of the targeted sites. Sediment from the targeted sites was, however, tested for toxicity to C. tepperi (see Results: Chironomus tepperi growth and emergence tests).
Sediment and surface water samples were analysed using a 98 component agrochemical screen of insecticides, herbicides and fungicides. The screen included the APLR-approved insecticides. These insecticides were tested with the limits of reporting (LOR) listed in Appendix 3 for the samples collected at sites and times described in Appendix 1. A suite of heavy metals, nutrients, hydrocarbons and organic carbon were also determined by a private consulting company (ALS laboratories, Melbourne). Water physico-chemical measurements (conductivity, pH, dissolved oxygen [DO], temperature, alkalinity [also by ALS], turbidity) also determined. Organic carbon, organic matter and total nitrogen were also determined for samples.
Chironomus tepperi growth and emergence tests
Sediments collected in the pesticide survey were tested for toxicity to C. tepperi larvae. Chironomus tepperi used in the present study originated from temporary ponds at Yanco Agricultural Institute in New South Wales (Choung et al., 2010). The culture was maintained in aquaria containing ethanol-sterilised tissue paper in artificial water made from a modified version of Martins solution (Martin et al., 1980) (reverse osmosis water with 0.12 mM NaHCO3, 0.068 mM CaCl2, 0.083 mM MgSO4, 0.86 mM NaCl, 0.015 mM KH2PO4, 0.089 mM MgCl2 and 0.1% (w/v) iron) at 21°C ± 1°C and minimum 65% relative humidity and a 16:8 h light:dark cycle photoperiod. The culture was fed slurry of ground Tetramin® in artificial water on alternate days after hatching. This mixture was also used in subsequent experiments.
The methods used to determine survival, sub-lethal acute and chronic effects were modified from the OECD guidelines (2004) and Stevens et al. (1993). Growth and emergence tests were carried out between September 2010 and August 2011.
For the growth assay, 10 five day old C. tepperi larvae were randomly added to beakers containing ca. 140 g (wet weight) sieved sediment and 200 mL of artificial water, with a minimum of three replicate beakers per sediment. After five days incubation at 21 °C (16:8 light:dark cycle) and prior to pupation, surviving larvae were removed from the sediment by sieving through a 125 μm sieve and body length was measured. Survival was also determined. Electrical conductivity, pH, dissolved oxygen and ammonia concentrations of the overlying water were measured at each water renewal during and at the end of the test. Artificial water was renewed every second day and larvae were given food at each water change.
For the emergence assay, ten 5 day old midge larvae were added to beakers containing ca. 140 g (wet weight) of sieved sediment and 200 mL artificial water, with a minimum of three replicate beakers per treatment. Beakers were incubated for 12-15 days (until laboratory control adult emergence reached at least 80%, as this ensures the test is valid (OECD, 2004) and is comparable to other sediment toxicity tests) at 21°C (16:8 light:dark cycle). The number of emerging adult C. tepperi was counted daily. Electrical conductivity, pH, dissolved oxygen and ammonia concentrations were measured at each water renewal during and at the end of the test. Artificial water was renewed every 2nd day and larvae were given food at each water change.
Copper (Cu) reference toxicity tests were also run for C. tepperi larvae from the same cultures used in the whole sediment toxicity testing, to assess whether the cultures were of appropriate sensitivity. Copper was used in this case as the survival response to this chemical had previously been characterised in the CAPIM laboratory for second instar C. tepperi larvae and is also used as a reference toxicant in routine sediment toxicity tests (Hai Doan, Commonwealth Scientific and Industrial Research Organisation [CSIRO], personal communication).
Statistical analyses were performed using the statistical software package Minitab® (statistical software release 15; Minitab, State College, PA, USA). Survival data were arc-sine square root transformed prior to statistical analysis. A General Linear Model and Dunnetts post-hoc tests were used to test for differences in growth and survival of C. tepperi between sites and the laboratory control. Differences between sites in the post-hoc tests were considered significantly different if p < 0.05. In all tests, data were checked to ensure that they conformed to the assumption of homogeneity of variance between groups and normal distribution of residuals.
Chironomid larval deformities
Chironomid larvae collected from any sites found to be contaminated with APLR-approved pesticides were analysed for morphological abnormalities. 3rd and 4th instar head capsules of genus Chironomus and Polypedilum were slide-mounted in Hoyers fixative, and observed under a compound microscope at 40-100 × magnification. Head capsules were observed for morphological abnormalities, following Warwick (1990). Structures examined were the antenna, mentum, ephypharangeal pectin and mandibles. Samples were analysed from sites where ≥ 15 larvae were collected. As these animals were collected from on site, this bioassay was considered a test of both sediment and surface water toxicity.
Statistical analyses were conducted in the software package R (www.r-project.org, version 2.10.1). Differences in incidence of deformities in Chironomus between treatment sites and a relatively unpolluted control site, Ferguson Paddock, Hurstbridge, Victoria, were tested using a chi-squared test. A chi-squared test was also conducted to test for differences in Polypedilum larvae collected from an unpolluted reference site, Glynns Wetland, Warrandyte, Victoria. Differences were considered statistically significant where p < .05.
Snail and amphipod in situ and laboratory mortality test trials
Cages containing freshwater invertebrates (snails and amphipods) were deployed in three monitoring sites (136 [Bendigo Creek, Huntly], 616 [Avoca River, Natte Yallock] and 622 [Campaspe River, Axedale]), as well as a field reference site (Glynns Wetland, Warrandyte, Victoria or Deep Creek, Bulla, Victoria) and a laboratory control, for seven days starting 11th November 2010 (in the case of the pre-spray occasion) and 10th March 2011 (in the case of the post-spray occasion). Cages deployed at a reference site and in the laboratory under controlled conditions were included to account for site habitat and water physico-chemical differences. Half the cages at each site contained site sediment and half were water-only exposures. In addition, post-spray sediment and water from the same three sites was transported to the laboratory for 7-d exposures of snails and amphipods to sediment + site water and site wateronly treatments under controlled conditions.
Snails (P. antipodarum) and amphipods (A. subtenuis) were collected from relatively noncontaminated streams in Victoria and held in the laboratory for no more than 48 h before field deployment in the cages or exposure in laboratory bioassays. Cages were constructed from clear polypropylene screw top containers (500 ml) with 500 μm nylon mesh covering side windows (40 mm × 50 mm) to allow water and oxygen flow-through.
Of the six cages deployed at each site, three replicate cages contained sieved (63 μm) sediment (0.5 cm depth layer) and three replicate cages did not have sediment (i.e. were water only exposure). All cages had a 30 mm x 30 mm piece of cotton gauze as artificial substrate and 300 mg of crushed fish food (Tetramin®). Twenty adult amphipods and 20 adult snails were randomly assigned to 50 mL tubes of culture media and transported to the deployment sites without mortality. Animals were randomly added to each cage on site immediately prior to cage deployment.
Physico-chemical measurements (pH, DO, conductivity [EC] and temperature) were taken for water at the time of cage deployment and retrieval. After one week, cages were retrieved and snail and amphipod survival was recorded. Surviving invertebrates were stored in liquid nitrogen for possible analysis of biomarkers at a future date.
For the laboratory-based experiments, sediment (140 g wet weight, 63 μm sieved) that had been collected from Sites 136, 616, 622 as well as reference sites were added to three replicate glass beakers (600 ml) and site water added (190 ml). A further three replicate beakers contained site water only. Laboratory controls consisted of reference site sediment with a standard artificial medium as the overlying water. Food (300 mg Tetramin®) and a 30 mm x 30 mm piece of cotton gauze acting as artificial substrate for the amphipods were added to each beaker. Fifteen adult snails and amphipods were randomly assigned to each beaker and all treatments were maintained under controlled conditions (16:8h light:dark; gentle aeration; 21 °C) for seven days. On day seven, physico-chemical parameters were measured and invertebrates were processed as for the in situ cage exposures.
To test for differences in invertebrate survival with site and water only versus water + sediment exposures, survival data was arcsine square root transformed to comply with homogeneity of variance and normality assumptions. A two-way analysis of variance (ANOVA) was performed using Minitab version 16 software with site and presence/absence of sediment as the two independent factors. Where statistically significant differences (p < 0.05) between sites were found, a Tukey's all pairwise multiple comparisons post hoc analysis was performed. A Dunnett's test was used to detect changes in survival between the reference and Sites 136, 616, 622. In the case of the pre-spray occasion, the sites were compared to the laboratory control because the reference site was affected by low DO. This was followed by one-way ANOVAs to detect differences in survival between pre-, post-spray and laboratory exposures within each site.
Determination of impacted sites
To determine whether sites were impacted by APLR-approved pesticides, a decision tree (Figure 4) was followed. This process considered data from all relevant analyses, and assigned each sample a likelihood of being impacted. Each sample was allocated one of four result categories: "very likely", "likely", "possibly" or "unlikely impacted" by the APLR. The lack of more definitive language can be considered acknowledgement of the fact that no environmental screening program can determine the presence of all possible contaminants. The possibility of unscreened chemicals being present must hence be considered, as must that that contaminants may be present below instrumental detection limits.
For chironomid, snail and amphipod experiments, a sample was considered "toxic" if any of the endpoints assessed yielded a toxic result relative to the unpolluted control. If a sample yielded a toxic result and a positive test for presence of any of any APLR-approved pesticide, the DPI Chemical Standards Field Services (CSFS) Team conducted an investigation of properties in the region surrounding the site. This investigation aimed to assess whether the detected pesticide had in fact been used in the APLR, or for pests other than locusts. If the latter were the case, then the sample was deemed "unlikely-imapcted" by the APLR. The presence of other toxicants, namely: non APLR-approved biocides, salinity, heavy metals, hydrocarbons and nutrients were systematically considered when assigning likelihoods of impacts to samples. If no potential toxicants were detected, yet toxicity was observed, it was assumed that toxicity was caused by an agent not screened for in the chemical analyses. Such samples were deemed "unlikely impacted" as all APLR-approved pesticides were part of the chemical screen. However, this does carry the assumption that concentrations below the limits of detection for these chemicals are sufficiently low as to not be toxic to monitored organisms.
Sites that yielded a "toxic" result for any post-spray sampling, but also for the pre-spray sampling, were deemed "unlikely" to be impacted. Many sites sampled only in the third sampling round (i.e. in south-western regions where locusts unpredictably migrated to) lacked a corresponding pre-spray sample. While these samples were still able to legitimately be classified as "toxic" or "not toxic" by comparison with the unpolluted control sample (ANZECC/AMRCANZ, 2000),the lack of a temporal reference sample lessened confidence in attributing toxicity to either APLR-approved pesticides or other causative factors. A less structured approached was adopted for the fish and the macroinvertebrate community monitoring. Where impacts were detected, evidence would be considered (e.g. observed impairment in organisms, presence of APLR-approved pesticides [where analysed], APLR activity in vicinity, other potential stresses such as flooding in area etc) in determining whether a site could reasonably deemed "possibly" or "unlikely impacted". Site 685 (Lake Koorlong) was monitored for fish impacts, but not for presence of chemical contaminants.
Only chemical results for APLR-approved pesticides are presented here. Three incidences of APLR-approved pesticide contamination in sediments were determined (see Table 4) and seven incidences in surface waters (see Table 5). Amber glass bottles containing surface water samples from Sites 137 (Wimmera River, Riverside), 139 (Wimmera River, Joel) and 626 (Kurrayah Swamp, Edenhope) sampled in the post spray round were broken in commute to the laboratory. These samples could therefore not be analysed for presence of biocides. See Appendix 4 and Appendix 5 for relevant levels of organic carbon, organic matter and total nitrogen.
|Sample Round||Site Code||Name||Locality||Sample Date||Insecticide||Concn (μg/L)||LOR (μg/L)|
Chironomus tepperi bioassays for sediments APLR-approved pesticides
Only results for sediments determined to be contaminated with APLR-approved pesticides are presented. Namely: Sites 130 (Tahbilk Wetland, Nagambie), 604 (Lake Hawthon, Mildura) and 659 (Lake Wongan).
At the end of each growth and emergence test the pH and DO values measured in the laboratory controls were pH 6.9 and 67.5% DO for growth, and pH 6.4 and 72.4% DO for emergence tests. These values satisfy the water quality criteria for test validity (OECD, 2004). pH and DO (% saturation) were generally similar across the sites and between tests, ranging between pH 7.32 to 8.5 and above 65% DO, except for Site 659 where DO was consistently between 40 and 60%. There were differences in the EC in the overlying water, in that Sites 604 and 659 had higher EC than Site 130 and the laboratory control sediment.
Survival and growth were endpoints used to determine the acute effects of contaminants in sediment. Survival in laboratory control beakers was consistently above 90% (ranging between 93-99%) following the five day exposure, this satisfies the test validity criteria set out by the OECD (2004). If survival in controls fell below 80%, then that batch of tests were repeated (OECD, 2004). Survival was high at all sites in the pre-spray sample, there was no significant difference in survival between field sites and the control (F (2,25) = 0.49, p = 0.62) at Site 130 at all sampling times, and at Site 659 in the third round sample (Figure 5). There were significant differences in larval survival in sediment collected from Site 604 in both post-spray (F (2,28) = 682, p = 0.000) and in the third round samples (F (3,24) = 12.01, p = 0.000) compared to laboratory control survival.
As the growth tests were carried out over a number of batches, the growth results were normalized to the control larvae, so are presented as percentage difference to controls (Figure 6). For statistical analysis each batch was analysed separately to compare with controls from that particular batch. There were significant reductions in larval growth from Sites 130 and 604 sediment compared to control larvae in their pre-spray samples (p = 0.000). Larvae exposed to post-spray and third round Site 130 sediment were not significantly different in length to control larvae. Larvae exposed to sediment collected from Site 659 did not show any adverse impacts on growth (p = 0.146) and, as this site was only sampled in the third round, a comparison with pre-spray results is not possible. As there was no survival following exposure to the post-spray sediment collected from Site 604, larvae lengths were not measured. Larvae exposed to Site 604 third round sediment were significantly smaller in length than controls (p = 0.017) but not significantly different to larvae exposed to the pre-spray sample (p > 0.05).
Emergence tests provide information about longer term chronic effects of exposure to contaminated sediment upon test organisms. They are a more sensitive test than the survival and growth test (Marinkovic et al., 2011). Sites which had emergence below 80% laboratory control were classified as 'marginally impacted', and below 60% were classified as 'severely impacted' (Anu Kumar, [CSIRO], personal communication) (Figure 7). Emergence tests conducted on pre-spray sediment from Site 130 showed a reduction in emergence relative to laboratory control, however post-spray and third round tests showed a significant improvement, and emergence was not significantly different to the control in either case. The emergence tests revealed the third round Site 604 sediment to be more toxic than pre-spray sediment. An emergence test was not conducted on the post-spray sediment collected from this site, as there was no survival in the five day survival and growth test.
Analysis of contaminated site-collected chironomids for deformities
Sufficient chironomid larvae were collected for morphological assessment for three of the 11 incidents of APLR-approved pesticide contamination. These were Site 150 (Lake Buninjon, Calvert) post-spray, Site 659 (Lake Wongan) third-round and Site 662 (Green Hill Lake, Ararat) third round. Forty-seven Chironomus larvae were collected from Site 150, 31 Chironomus larvae from Site 659, and 15 Polypedilum larvae from Site 662. As deformities were observed predominantly in antennae, almost to the exclusion of all other structures, only incidence of antennal deformities were statistically compared to control sites. Antennal deformities observed were reduced antennal blades, and misshapen antennal segments.
For Chironomus larvae, a chi-squared test showed that Site 150 did not have significantly elevated antennal deformities (p > 0.05), whereas Site 659 did (p < 0.05).
For Polypedilum larvae, a chi-squared test showed that Site 662 did not have a significantly elevated incidence of deformities relative to the control sample (p > .05).
Snail and amphipod bioassay trials
Physico-chemical parameters of conductivity (EC), pH, dissolved oxygen (DO) and water temperature measured at the time of deployment and retrieval of the in situ cages varied across sites and between pre- and post-spray occasions (see Table 6). The lowest to highest mean values measured for each parameter were EC:84.5 to 3,160 μS/cm, pH 6.0 to 8.1, DO:11 to 106 %, and temperature: 13 to 21 °C. At Site 616 (Avoca River, Natte Yallock), dissolved oxygen was at the lower threshold of physiological tolerance for most invertebrates on both pre- and post-spray occasions and at the control site (Glynns Wetland) on the pre-spray occasion. The variability in DO and temperature was mitigated when testing was conducted with site specific water and sediment in the laboratory and these parameters stabilised at >80% and 21 °C respectively (see Table 6).
|In situ cages|
|Laboratory water-only exposures|
|Laboratory water+ sediment exposures|
The APLR-approved pesticide fipronil (and its breakdown products: fipronil sulfide, fipronil sulfone and fipronil desulfinyl) were detected (each at 0.001 µg/L) at one of the three monitoring sites (Site 136 [Bendigo Creek, Huntly]) on the post-spray occasion in water. Arsenic measured in sediment at Site 136 (279 mg/kg) exceeded the ANZECC/ ARMCANZ (2000) ISQG high sediment quality trigger value (70 mg/kg).
No significant differences were found in amphipod survival between cages with and without sediment, therefore these treatments were pooled in order to test for differences between sites and pre and post spray occasions. Pre-spray caged amphipod survival at all sites was either not different or significantly higher than in the caged laboratory control. Post-spray survival was significantly lower in the caged laboratory control (67 ± 9 % survival, mean ± SD), Site 616 (0.83 ± 3 % survival, mean ± SD) and Site 622 (Campaspe River, Axedale) (77 ± 20% survival, mean ± SD) when compared to the reference. Significant reductions in caged amphipod survival between pre- and post-spray deployments at a site occurred on the pre-spray occasion at the reference site (37 ± 6% survival, mean ± SD) and on the post-spray occasion at Sites 616 and 622 (Figure 10). As previously noted, the DO at the pre-spray reference site (13 ± 20% DO, mean ± SD) and the post-spray deployment at Site 616 (11 ± 2% DO, mean ± SD) was at the lower limits of physiological tolerance and likely explained the observed effects on survival.
Survival of amphipods increased to 99% at Site 616 when the artefact of low DO was removed by exposure of amphipods to the same sediment and water under controlled laboratory conditions with aeration provided (Figure 10). Site 622 also had higher survival under aerated laboratory conditions (100 ± 0% survival, mean ± SD) than under caged conditions in the field (77 ± 20% survival, mean ± SD) on the post-spray occasion, although DO was not physiologically challenging at this site (64 ± 8% DO, mean ± SD). There were no significant differences in survival of amphipods between the reference and other sites when amphipods were exposed to post-spray samples under laboratory conditions.
No significant differences were found in snail survival between those cages or laboratory exposures with and without sediment, therefore these treatments were pooled in order to test for differences between sites and pre and post spray occasions. Across all sites in all field and in the laboratory deployment occasions, snail survival ranged between 82 ± 20% and 100 ±20 % (mean ± SD) (Figure 11). No significant decrease in survival was detected (p > 0.05) relative to the reference or other sites.
Threatened fish monitoring
A total of 492 Murray hardyhead were recorded during APLR monitoring (Table 7).
|Location (Date sampled)|
|613. Woorinen North Lake|
|614. Round Lake|
|612. Cardross Lakes Basin 1|
|685. Lake Koorlong|
|Double Wing Fyke Netting||0||471||18||1|
At Site 613 (Woorinen North Lake) no Murray hardyhead were captured despite a more intensive sampling regime (utilising all three sampling techniques) being employed here. Approximately 10,000 mosquitofish (Gambusia holbrooki) and eight carp (Cyprinus carpio) were captured here (Stoessel, 2011).
At Site 614 (Round Lake, Lake Boga), a total of 471 Murray hardyhead consisting of adults and juveniles were captured with double winged fyke nets. 18 adult Murray hardyhead were captured with double winged fyke nets from Site 612 (Cardross Lakes Basin1). Twenty-two unspecked hardyhead were also captured from the double winged fyke nets here.
At Site 685 (Lake Koorlong), three adult Murray hardyhead were captured using boat electrofishing and fyke netting techniques. Over 20,000 mosquitofish were also collected from this lake (Stoessel, 2011).
The 20 sites sampled in the "targeted" part of the program were in relatively poor condition when assessed against the single season objectives for the indicators used in SEPP (Waters of Victoria) (see Appendix 6 for all scores). Of the 40 samples collected (two from each site), only four met the objectives. The protocol specified in the policy is that both samples collected from a stream site need to meet the objectives. On this basis all stream locations failed this test prior to any spraying. This result is not unexpected as the majority of streams in the northwestern catchments of Victoria have failed in recent years, due to poor water quality and drought impacting on the quantity of water present.
The Sustainable Rivers Audit had up to 35 sites in each these catchments: Broken, Campaspe, Avoca and Wimmera. The majority of results are based on combined seasonal data (spring and autumn) as required in the SEPP. Some sites in each catchment have been assessed against single season objectives. Results for each sample are summarised in Table 8. These results show a poor rate of meeting the policy objectives, ranging from 28% in the Broken to 7% in the Avoca.
|Catchment||Total number of samples||% Failed in 10/11||% Passed in 10/11||% Passed in 08/09|
Investigation of APLR activity in areas where toxicity was observed for samples
Three incidences of APLR-approved pesticide contamination were found to be associated with toxicity in test organisms. These were Site 604 (Lake Hawthorn) post-spray, Site 604 third round and Site 659 (Lake Wongan) third round. Chlorpyrifos-contaminated Site 604 and Site 659 sediments induced toxicity in laboratory C. tepperi experiments, and Site 659 sediment was also associated with elevated deformities in field-collected Polypedilum larvae.
As per the decision-tree (Figure 4), properties in the vicinity of sites found to be yielding toxic samples were investigated by the DPI CSFS to determine whether significant APLR activity, potentially leading to aquatic contamination, had occurred in the area.
This was found not to be the case for either site. Minimal APLR activity had occurred in the Site 604 region, and chlorpyrifos use was found to be in the main for treatment of mealybug, oriental fruit moth and light brown apple moth infestations in nearby vineyards. No APLR activity was found to have occurred in the vicinity of Site 659, and chlorpyrifos use here was found to be instead associated with lucrene flea and aphid control in surrounding canola crops (Alan Roberts, DPI, personal communication).
Determination of sites impacted by APLR
All sites were determined to be "unlikely impacted" by the APLR. This excludes sites assessed only for macroinvetebrate community effects, upon which no conclusions were drawn.
APLR-approved pesticides were not found above the limits of detection at the vast majority of these 81 sites assessed for toxicity to test organisms.
Of the 11 incidences of contamination in post-spray and third round sampling, APLR impacts were ruled out due to either the lack of a toxic result, the lack of increased post-spray toxicity (relative to post-spray sample) or a DPI CSFS investigation finding that the contamination observed was a result of non-locust pest control (See Table 9 and Table 10). Although the Site 638 (Joyces Creek, Strathlea) surface water sample could not be toxicologically assessed (as insufficient chironomid larvae were collected for deformity analysis, and it not being part of the snail and amphipod bioassay trials), the same concentration of carbaryl was determined not to be toxic for Green Hill Lake. For this reason, it was concluded that the Site 638 was "unlikely impacted" by the APLR.
The third round incidence of surface water contamination (fipronil, fipronil sulfide and diazinon) at Site 136 (Bendigo Creek, Huntly) could also not be toxicologically assessed due to a lack of chironomid larvae collected and this round not being part of cage bioassay trials. As concentrations of fipronil (0.006 μg/L) and diazinon (0.02 μg/L) were considerably lower than their literature LC50 values to C. tepperi (0.43 μg/L and 35.5 μg/L respectively), these were thought somewhat unlikely to cause at least a mortality impact in this test organism. For this reason, the third round Site 136 surface water sample was tentatively deemed "unlikely impacted" by APLR. It should be noted that sublethal effects or lack thereof cannot be inferred in this instance. There are no literature toxicological values for fipronil breakdown products to C. tepperi. All four sites where Murray hardyhead populations were monitored were deemed "unlikely impacted". Three of the populations appeared relatively healthy, and a paucity of animals captured in Site 613 (Woorinen North Lake) was attributed to causes other than the APLR. The "unlikely impacted" classification was primarily due to a lack of APLR-approved pesticides detected in the lake, but also other potential causes were observed (see Discussion). Site 685 (Lake Koorlong) was not part of the pesticide survey so was not assessed for presence of pesticides, but was deemed "unlikely impacted" due to the apparent health of its Murray hardyhead population.
No conclusions were drawn as to the likely impacts of APLR on sites based on macroinvertebrate monitoring results, due to potential confounding effects of heavy rains and flooding.
|Sample||Sampling Round||Endpoint||Endpoint assessed?||
More toxic than |
More toxic |
|Bioassay results||Site results|
|C. tepperi mortality||Yes||Yes||Yes||
No significant |
|C. tepperi growth||No||N/A||N/A||N/A||N/A|
|Snail mortality trial||No||N/A||N/A||N/A||N/A|
|C. tepperi mortality||Yes||Yes||Yes||No significant locust control in area||
|C. tepperi growth||Yes||Yes||Yes||No significant locust control in area||
|Yes||Yes||Yes||No significant locust control in area||
|Snail mortality trial||No||N/A||N/A||N/A||N/A|
|C. tepperi mortality||Yes||No||N/A||N/A||N/A|
|C. tepperi growth||Yes||No||N/A||N/A||N/A|
|Snail mortality trial||No||N/A||N/A||N/A||N/A|
|C. tepperi mortality||Yes||No||N/A||N/A||
|C. tepperi growth||Yes||No||N/A||N/A||
No locust control in|
No locust control in|
|Snail mortality trial||No||N/A||N/A||N/A||N/A|
|Amphipod mortality trial||No||N/A||N/A||N/A|
Determination of sites impacted by APLR
Of 81 study sites monitored for ecotoxicological effects in this study, none were determined to have been ecologically impacted by pesticides applied in the APLR. A variety of ecotoxicological endpoints were employed to maximise the likelihood of observing any possible toxic effects. Importantly, sites were selected that were thought most likely to receive agricultural runoff in locust risk areas, in order to maximise the likelihood of detecting any contamination. The use of ecotoxicological assays in this study has been a demonstrable improvement on traditional monitoring programs that are based soley on chemical analyses. These traditional programs have a limited capacity to infer toxicity, or lack thereof, of contamination.
Although the detection of carbaryl in post-spray surface water from Site 638 (Joyces Creek, Strathlea) was unable to be toxicologically assessed (due to the low number of chironomid larvae collected for deformity analysis and cage bioassay trials not being conducted at this site), the same concentration of carbaryl was found to be non-toxic for Site 662 (Green Hill Lake).
The detection of fipronil, fipronil sulfide and diazinon in third round Site 136 (Bendigo Creek, Huntly) surface water was unable to be toxicologically assessed for the same reasons as for Site 638. However, it can be at least said that the concentrations observed were somewhat unlikely to exert a mortality effect. The determined concentrations of fipronil and diazinon (0.006 and 0.02 μg/l, respectively) were both orders of magnitude lower than the literature LC50 for C. tepperi (0.43 and 35.5 μg/L respectively).
The detection of fipronil in pre-spray surface water at Site 615 (Lake Cooper, Corop) was deemed not to be associated with the APLR, as it occurred prior to the likely commencement of spraying. Furthermore, no fipronil was detected in post-spray sampling at Site 615.
The possibility of collecting insufficient chironomid larvae for deformity analysis has meant that in some cases, APLR-approved pesticides detected in surface waters has gone unassessed for toxicological effects. It is therefore apparent that in future monitoring events, bioassays for surface waters, such as the snail and amphipod bioassays trialled in the experiment, are desirable.
Incidence of APLR-approved pesticide contamination
Each of the detected APLR-approved pesticides reported is approved for a range of insect pests on a range of agricultural crops; and fipronil is also registered as a termiticide for control in domestic dwellings. The chlorpyrifos detected in the Site 604 (Lake Hawthorn, Mildura) samples was associated with presence of another pesticide, prothiofos in a similar range of concentrations. Prothiofos is an organophosphorus insecticide registered in Victoria only for control of caterpillars in brassica vegetable crops and mealybug in table grapes and pears. There was a significant infestation of mealybug in table grapes through the Mildura area in summer 2010-11. Although not registered for APLR control it is likely that prothiofos would control APL in a similar efficacy to fenitrothion, chlorpyrifos, diazinon and malathion. Chlorpyrifos, in contrast to prothiofos, has in excess of ninety different product registrations for insect control on a range of crops, domestic situations and as a termiticide, and is registered for control of mealybugs in grapes.
Amber glass bottles containing post-spray surface waters for Sites 137 (Wimmera River, Riverside), 139 (Wimmera River, Joel) and 626 (Kurrayah Swam, Edenhope) were broken in transit, meaning these samples could not be analysed for presence of APLR-approved or other biocides. It cannot be commented on as to whether these water samples were contaminated with APLR-approved pesticides, other than to say it is unlikely, given the low overall detection rate of these pesticides. Furthermore, all three of these sites were follow-up sampled in the third round, and APLR-approved pesticides were not detected in surface waters from any of these sites.
Chironomus tepperi sediment bioassays
Of the sediment samples contaminated with APLR-approved pesticides, only Site 604 (Lake Hawthorn, Mildura) sediment showed all three of C. tepperi survival, growth and emergence effects. Post-spray sediment was highly acutely toxic to C. tepperi and resulted in 100% mortality after a five day exposure, hence no subsequent tests were carried out on this sediment. The third round Site 604 sediment samples showed a significant increase in survival (57%), but this was still significantly lower than laboratory control survival. Growth and emergence from the third round samples were significantly lower than controls, and emergence only was significantly lower than pre-spray-exposed larvae. There were a number of other chemicals detected in the sediment in the post-spray and third round samples compared to the pre-spray sediment that may also have contributed to the observed effects on C. tepperi larvae.
Site 659 (Lake Wongan), sampled only in the third round, showed only emergence impacts. Chlorpyrifos (an organophosphate pesticide) was detected in sediment collected from both Sites 604 and 659. Chlorpyrifos concentrations at Site 604 were higher than at Site 659 (28 and 22 µg/kg sediment at Site 604 for post-spray and third round, respectively, compared to 10 µg /kg at Site 659). Direct comparison between Site 604 and 659 results is impossible however, as several other biocides, including prothiofos, were detected in Site 604 sediments. Hence no toxic result for 604 can be unequivocally linked to chlorpyrifos contamination. Nonetheless, chlorpyrifos is highly acutely toxic to chironomids, with a reported LC50 for C. tepperi in water of 1.3 µg/L (Stevens, 1992). There is limited information about the effects of sediment-bound chlorpyrifos on chironomid life history parameters published in the literature; the majority provide information only about the acute toxicity in water exposures. Callaghan et al (2001) did however find male emergence was delayed at when Chironomus riparius were exposed to sediment concentrations of 0.1 mg /kg of chlorpyrifos.
Cyhalothrin was detected in the post-spray sediment from Site 130 (Tahbilk Wetland, Nagambie), and has been shown to be toxic to the chironomid Chironomus tentans (Bouldin et al., 2005) However, our results suggest that the concentration present is too low to be toxic to C. tepperi larvae, given the lack of toxic results observed for all endpoints. Sediment collected from this site during the third round was not toxic and cyhalothrin was not detected on this occasion.
Emergence tests are useful in sediment toxicity tests as they can incorporate long term effects that may be exerted at later stages of development (Marinkovic et al., 2011) or more subtle effects on life cycle responses (Paumen et al., 2008). If emergence tests on Site 659 sediment had not been conducted survival and growth data would infer that this sediment was not toxic to C. tepperi, therefore it is important that sediment toxicity tests include long-term as well as short-term tests. This was also observed in a biomonitoring study by Kellar et al. (2011), where exposure to contaminated sediment had no effect on C. tepperi survival but affected growth and emergence. The advantage of conducting the short term growth test is that the acute toxicity of the sediment can be established in a relatively short time frame.
Chironomid larval deformities
Of the 11 incidences of APLR-approved pesticide contamination, sufficient chironomid larvae for deformity analysis were collected in three. Chironomus larvae collected from Site 659 showed a significant elevation in antennal deformities. These deformities are possibly attributable to chlorpyrifos, as this was the only contaminant detected and is known be toxic to chironomids (Callaghan et al., 2001). Though nickel concentrations in sediment exceeded the ISQG low concentration (ANZECC/AMRCANZ, 2000), it naturally occurs at these concentrations in Victorian basalts in this region (Pettigrove, 2006) and is unlikely to have caused an increased incidence of mouthpart deformities. Pesticide concentrations were not associated with increased deformities in Chironomus collected from Site 150 (Lake Buninjon, Calvert), nor in Polypedilum from Site 662 (Green Hill Lake, Ararat).
While these deformity results are indicative of a toxic effect either occurring or not occurring, a mechanistic explanation for the inducement of deformities has not yet been determined. Results are therefore correlative (Diggins and Stewart, 1998) and cannot unequivocally determine a specific causative agent.
Snail and amphipod bioassay trials
There were no significant differences in survival of amphipods and snails in the field cages or the laboratory exposures between those treatments with and without sediment, suggesting that any observed responses were not due to the presence of sediment. Suspended particulate material within the water may still have contributed to responses. Differences in survival of amphipods caged at three potential pesticide spray sites relative to the caged laboratory control (in the case of the pre-spray occasion) or reference site (in the case of the post-spray occasion) were likely driven primarily by low DO rather than exposure to contaminants associated with either the water or sediment. This was supported by the improved survival of amphipods when adequate DO was supplied under laboratory conditions in the case of Sites 616 (Avoca River, Natte Yallock) and 622 (Green Hill Lake, Ararat) where there were survival differences between pre and post-spray cage deployments.
Snails proved to be robust to low dissolved oxygen levels at the field sites. No significant effects on snail survival were observed in any experiment. Snails and amphipods exhibit median lethal toxicity to salinity (LC50 ≥ 12,000 and 52,000 µS/cm, respectively (Kefford et al., 2003); 4 to 17 times higher than the maximum measured site conductivity (3,120 µS/cm ) (Table 6) therefore, salinity was not responsible for observed effects on amphipod survival. Overall, based on the responses of caged and laboratory exposed amphipods and snails, there were no acutely toxic effects on survival due to APLR-approved pesticides, in either water or sediment at Sites 136 (Bendigo Creek, Huntly), 616 and 622.
Threatened fish monitoring
Monitoring of threatened fish was only conducted on Murray hardyhead. Threatened species not monitored in this study (silver perch, unspecked hardyhead, trout cod, Murray cod, golden perch, Macquarie perch and catfish) were not assessed due to various environmental and operational issues, namely:
- Unprecedented and continued extensive flooding of sampling areas.
- Inability to access sprayed areas within timeframes required for meaningful results to be obtained.
- APL spraying did not proceed in some target locations.
As such, we are unable to conclude whether or not the locust control program impacted on these threatened fish species. Additionally, some threatened species populations also experienced a blackwater event. Blackwater events occur when flood waters inundate floodplains or dry river channels, and carbon compounds are leached from accumulated river red gum (Eucalyptus camaldulensis) leaf litter (including, leaves, twigs and bark) resulting in high levels of dissolved organic carbon, which can often lead to hypoxia (extremely low levels of dissolved oxygen) within the stream (King et al., 2011). Blackwater may also be the result of agricultural runoff. Blackwater may have impacted fish populations in areas where it occurred, some of which were within the plague locust risk zone.
Two of the four Murray hardyhead populations Site 614 [Round Lake, Lake Boga] and 612 [Cardoss Lakes Basin 1] surveyed in the current study appeared to be in good health.
The lack of Murray hardyhead captured from Site 613 (Woorinen North Lake) could indicate that this species has not recovered from the crash in the population that had occurred here between Spring 2009 and Autumn 2010 (Stoessel, 2010); prior to the APLR. Additionally, no APLR-approved pesticides were detected at Site 613, therefore APLR was deemed unlikely to have been responsible for the lack of captured Murray hardyhead. Competitive pressure from the mosquitofish population here may also be contributing to the lack of Murray hardyhead. Murray hardyhead exhibit a major niche overlap with mosquitofish with respect to both habitat (eggs, larval, juvenile and adult) and dietary (larval, juvenile and adult) requirements. Potential impacts are: predation of eggs and or larvae, competition for food or habitat and aggression such as fin nipping (Mac Donald and Tonkin, 2008).
Site 685 (Lake Koorlong) also showed a very low abundance of Murray hardyhead, while abundance of the introduced mosquitofish was very high. Site 685 was not part of the pesticide survey, though fish survey results suggest a healthy mosquitofish population, and hence little likelihood of toxic contamination at least for this species. The mosquitofish population may also be competing with Murray hardyhead here, possibly contributing to low Murray hardyhead numbers.
The population of Murray hardyhead in Site 614 appears to be healthy with no evidence of impact by APLR-approved pesticides. The pesticide survey also found no APLR-approved pesticides above the limits of detection in Site 614 samples. The presence of Murray hardyhead juveniles (<20 mm) indicates that there has been successful recruitment from the spring 2010/summer 2011 cohort, during the APLR. This suggests that pesticide application for the control of locusts in the Site 614 area did not negatively impact the Site 614 Murray hardyhead population.
Although adult Murray hardyhead were captured at Site 612, no juveniles were detected. No APLR-approved pesticides were detected in Site 612 samples, indicating that an APLR effect on the juveniles of this species is unlikely. The lack of juveniles within this population is more likely attributable to this species having not yet spawned prior to our surveys. Previous research has shown that Murray hardyhead in the Site 612 have a long breeding season between late September and April/May (Stoessel, 2010).
The Silversides family (of which hardyhead are part) may be sensitive to Metarhizium anisopliae spores, especially at the larval and juvenile stages (Genthner and Middaugh, 1995). At the time of writing it is unknown whether Metarhizium was used in the vicinity of Sites 685 or 613.
Although not part of the original fish monitoring study design, ARI also responded to two events occurring during the period of locust control. Twelve common yabbies (Cherax destructor) were found dead in a drainage outlet that runs into Site 604 (Lake Hawthorn, Mildura) on 5th October 2010. This coincided with the post-spray sampling for the pesticide survey, and the aforementioned chlorpyrifos contamination (see Results: Investigation of APLR activity in areas where bioassays on test organisms yielded toxic results).
The second event requiring ARI response was a Murray cod (Maccullochella peelii peelii) death and the disappearance of shrimp from the Pyramid Creek downstream of the Murrabit Road bridge being reported around the 24th November 2010. This was during a period of intensive APL control throughout the district (Rob OfBrien, DPI Kerang, Victoria personal communication). Staff from ARI working on the present study became aware of this report on Monday the 29th November. Shortly after this report, significant flooding occurred in Pyramid Creek and thus a sediment sample was unable to be attained to confirm or refute pesticide contamination in this stream. There were also some reports of blackwater events occurring in the area, though evidence for this was largely anecdotal and conflicting (Ross Stanton, Goulburn Murray Water, personal communication). The cause of the Murray cod death and disappearance of shrimp was therefore unable to be determined.
Due to weather and flooding forcing the abandonment of field sampling and possibly confounding results, the macroinvertebrate monitoring program was unable to conclude whether or not the locust control program had impacted on monitored communities. Nonetheless, several improvements for future locust control monitoring programs are apparent. Had data been immediately available as to locust insecticide applications (either on public or private land) in any of the monitored areas, it is possible that macroinvertebrate sampling could potentially have occurred in the window of time subsequent to spraying but prior to heavy rains. Hence flooding could have been ruled out as a causal factor in any impacts observed. However, in the present study, field teams were only made aware of more general data, in the form of public land applications in the broader regions (e.g. North Central region).
Rather than relying on an opportunistic sampling design described above, an alternative is to work with a few landholders who will be applying the pesticide and establish a more controlled experimental situation. In this way a "Before-After Control-Impact" (BACI) (ANZECC/AMRCANZ, 2000) design could be employed. This would most likely assess the effectiveness of spray applications according to the recommended protocols; a more opportunistic approach is more likely to pick up where protocols are not followed and would act as a more effective audit of the program.
Future monitoring programs might also incorporate quantitative methods. The standard "Rapid Bioassesment" (RBA) methods employed in this study only give consideration to presence-absence data for macroinvertebrate families. Pesticide contamination may reduce the abundance of sensitive aquatic macroinvertebrate species, so abundance data may supplement RBA data and produce more informative results.
Given that some detections for APLR-approved pesticides were in lakes, future macroinvertebrate monitoring programs might also incorporate lake and wetland sites, as well as rivers and creeks. It is possible that heavy rains flushed contaminated sediments and surface waters to lakes at the bottom of catchments, and here lake and wetland macroinvertebrate monitoring could be conducted under standard EPA guidelines (2010).
In conclusion, this study found no evidence that the APLR caused ecologically toxic contamination of rivers and wetlands with pesticides. The relatively small number of samples contaminated with APLR-approved pesticides were either non-toxic to test organisms, or found to be unlikely contaminated due to the APLR. Due to weather impacting field surveys, it was unable to be determined whether the APLR impacted monitored macroinvertebrate communities. Locust plague numbers (and consequently pesticide use) were much lower in this plague event than would reasonably be expected in subsequent events (due to atypical weather conditions), so caution must be observed when making inferences for future control programs.
This study has proven invaluable in determining with a high degree of confidence whether or not contamination and ecological impacts have occurred as a result of the APLR. The authors conclude that a similar monitoring program, assessing both chemical and biological endpoints would be essential in determining any environmental impacts for future large-scale pest control events. As an outcome of this project, the Victorian community can be reasonably assured that APLR activities did not cause major aquatic environmental impacts in regional areas. The authors recommend considerable improvements with respect to reporting of pesticide applications in future locust plague events; in terms of types, volumes, locations and timing of applications. This data should be made available to biomonitoring programs immediately subsequent to any application event.
Recommendations for future locust control programs
- Better reporting of pesticide use in locust control. Currently, data for use of pesticides on private and public land is limited. Private land use data is only available for land managers who make rebate claims in the LIRS. As well as being limited,data was often delivered after environmental monitoring was conducted. This reduced the capacity to confidently determine whether observed pesticide detections were associated with application for the purpose of locust control. Crucially, data should be specific with regards to chemicals used, timing, area and volume of applications, and should be made available to biomonitoring projects (be they chemical, macroinvertebrate, fish or laboratory toxicity monitoring programs, or a combination of these) as soon after each application event as practicable.
- Maintenance of current strategies to minimise risk of entry of contaminants into waterways (e.g. restriction zones, use of Metarhizium, avoidance of spraying adults).
- Future macroinvertebrate community and fish population monitoring programs should include sediment and water pesticide analyses and toxicity testing for every macroinvertebrate sampling event; to better assess pollution as a causal factor in observed effects.
- Employment of additional bioassays to test toxicology of surface water contamination, such as the snail and amphipod methods trialled in this study.
This study was funded by the Department of Primary Industries Biosecurity Victoria Division. The authors would like to thank Rebecca Brown for assistance in maintenance of the chironomid cultures and also for running the sediment toxicity tests. The authors would also like to thank Daniel MacMahon, Cameron Amos, Claudette Kellar, Stephen Marshall, Matthew O'Brien, Rowan MacMahon, Valentina Colombo, Iain Ellis, Anthony Cablel and Daniel Stoessel for assistance with field work. Thanks also to Daniel MacMahon and Cameron Amos for additional laboratory assistance.
The authors would also like to thank Dr Sze Flett from ARI for assistance in administration of this project, and to Alan Roberts from DPI for conducting field pesticide use investigations. Thanks also to the North Central, Wimmera, Mallee and Goulburn Broken CMAs, and DPI Victoria for assistance with selection of field sites.
Thanks to Stephen Marshall for comments on an early draft of this report and Emily Thomson for formatting the document.
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Appendix 1: Study sites, sampling regime (1 = pre-spray sampling round, 2 = post-spray sampling round, 3= third sampling round) and endpoints assessed (C = chemical analysis, CT = C. tepperi toxicity, PA = snail mortality, A = amphipod mortality, CD = chironomid deformities, MHH = Murray hardyhead population effects)
|Site Code||Site||Location||Sampled in rounds||Endpoints assessed|
|640||Loddon River||Baringhup West||2||C,CT|
|637||Bet Bet Creek||Norwood||2||C,CT|
|643||Avoca River||Gowar East||2||C,CT|
|646||Avoca River||Gowar East||2||C,CT|
|632||Bet Bet Creek||Norwood||2||C,CT|
|636||Loddon River||Baringhup West||2||C,CT|
|650||Avon River||Marnoo East||2||C,CT|
|651||Avon River||Marnoo East||2||C,CT|
|663||Lake Lonsdale||Mount Dryden||3||C,CT|
|665||Tea Tree Lake||Kanagulk||3||C,CT|
|667||Konoong Wootong Reservoir||Konong Wootong||3||C,CT|
|652||Lake Goldsmith||Lake Goldsmith||3||C,CT|
|658||Lake Burrumbeet||Bo Peep||3||C,CT|
|659||Lake Wongan||Lake Wongan||3||C,CT,CD|
|660||Lake Bolac||Lake Bolac||3||C,CT|
|662||Green Hill Lake||Ararat||3||C,CT,CD|
|670||Hopkins River||Langi Logan||3||C,CT|
|681||Mt William Swamp||Willaura||3||C,CT|
|682||Lake Turangmoroke Inlet||Lake Bolac||3||C,CT|
|684||Slate Lake Inlet||Mount Emu||3||C,CT|
|616||Avoca River||Natte Yallock||1,2||C,CT,PA,A|
|129||Goulburn River||Kirwans Bridge||1,2||C,CT|
|606||Lake Buloke||Lake Buloke||1,2||C,CT|
|612||Cardross Lakes Basin 1||Cardross||1,2||C,CT,MHH|
|628||Lake Cognumbul||East of Awonga||1,2,3||C,CT|
|615||Lake Cooper||Corop/Lake Cooper||1,2||C,CT|
|630||Lake Cullen||Lake Cullen||1,2||C,CT|
|629||Little Lake Boort||Boort||1,2||C,CT|
|140||Mt William Creek||Barton||1,2||C,CT|
|614||Round Lake||Lake Boga||1,2||C,CT|
|613||Woorinen North Lake||Woorinen North||1,2||C,CT,MHH|
|136||Bendigo Creek||Huntly||1,2,3||C,CT, PA, A|
|130||Tahbilk Wetland||Tahbilk Winery||1,2,3||C,CT|
Appendix 2: Sites for EPA targeted macroinvertebrate monitoring
|EPA sitecode||Site name||Basin||Sample datetime||Latitude||Longitude||MGA zone||MGA easting||MGA northing||Habitats sampled|
|GGH||Bet Bet Creek @ Norwood||Loddon||20/10/10 10:57||36.9944||143.6420135||54||735116||5902478||Dual edge|
|GHN||Middle Creek d/s Saligaris Rd||Loddon||18/10/10 18:02||37.1592||143.9069214||54||758137||5883497||Dual edge|
|GHO||Joyces Creek d/s Hurns Rd||Loddon||18/10/10 14:59||37.1292||143.9618988||54||763124||5886678||Dual edge|
|GHP||Joyces Creek west of Captains Gully Rd||Loddon||18/10/10 13:11||37.1492||143.9625702||54||763114||5884455||Dual edge|
|GJW||Loddon River Opposite Okeefs Rd||Loddon||20/10/10 16:21||36.9431||143.9060059||54||758788||5907478||Dual edge|
|GLE||Bet Bet Creek 500m d/s of GGH||Loddon||20/10/10 10:00||36.9926||143.6484528||54||735695||5902657||Dual edge|
|GLF||Joyces Creek d/s of GHO||Loddon||18/10/10 15:01||37.1214||143.9620514||54||763164||5887535||Dual edge|
|GLG||Middle Creek d/s GHN||Loddon||18/10/10 17:50||37.1529||143.9035187||54||757856||5884201||Dual edge|
|GLH||Loddon River @ Barringhup West Rd||Loddon||20/10/10 14:58||36.9428||143.9353638||54||761405||5907435||Dual edge|
|GLI||Joyces Creek 500m u/s of GHP||Loddon||18/10/10 14:01||37.1533||143.9631653||54||763153||5884002||Dual edge|
|HGN||Cherrytree Creek @ Moyreisk||Avoca||20/10/10 12:53||36.9033||143.4326935||54||716744||5913076||Dual edge|
|HJG||Avoca River off Wedderburn Rd||Avoca||19/10/10 12:53||36.5504||143.3996277||54||714779||5952309||Dual edge|
|HJI||Fentons Creek @ Logan||Avoca||19/10/10 10:31||36.6218||143.489563||54||722624||5944183||Dual edge|
|HLA||Fentons Creek 500m d/s Wimmera Hwy||Avoca||19/10/10 10:38||36.6171||143.4909363||54||722761||5944702||Dual edge|
|HLB||Avoca River 1km u/s Wedderburn Rd||Avoca||19/10/10 12:51||36.5542||143.4098816||54||715685||5951860||Dual edge|
|HLD||Strathfillan Creek @ Archdale Rd||Avoca||21/10/10 11:54||36.7383||143.3369904||54||708663||5931601||Dual edge|
|HLE||Strathfillan Creek u/s Dunolly Rd||Avoca||19/10/10 0:00||36.726||143.34375||54||709300||5932951||Dual edge|
|HLF||Cherrytree Creek u/s of HGN||Avoca||20/10/10 12:04||36.9043||143.4307251||54||716565||5912969||Dual edge|
|IHM||Avon River @ Boyles Bridge||Wimmera||19/10/10 15:38||36.6406||142.9792175||54||676938||5943161||Dual edge|
|INE||Avon River u/s IHM||Wimmera||19/10/10 15:00||36.6419||142.9804993||54||677050||5943011||Dual edge|
Appendix 3: Limits of reporting (LOR) forAPLR approved insecticides
|Insecticide||Sediment LOR (μg/kg)||Surface water LOR (μg/L)|
Appendix 4: Organic carbon and organic matter levels for sediments contaminated with locust-registered pesticides
|Organic Carbon||Organic matter|
|Sampling Round||Site number||Body||Name||Locality||Sample date||g/100g||g/100g|
|Post?\spray||130||Wetland||Tahbilk Wetland||Mulberry Drive||27/01/11||2.1||3.8|
|3rd round||604||Wetland||Lake Hawthorn||Mildura||17/03/11||2.1||3.8|
|3rd round||659||Wetland||Lake Wongan||Lake Wongan||9/06/11||8.8||16.0|
Appendix 5: Total organic carbon (TOC) and Total Nitrogen levels for surface waters contaminated with locust-registered pesticides
|Sampling Round||Site Code||Body||Name||Locality||Sample Date||TOC mg/L C||Total Nitrogen mg/L|
|Post?\spray||662||Wetland||Green Hill Lake||Ararat||6/06/11||19.0||2.10|
Appendix 6: Macroinvertebrate results from the EPA "targeted" sites sampled.
|Site code||Replicate||Region||Habitat||OE5 0||Band||SIGNAL||Total Taxa Richness||EPT Taxa||Sample Attained Objectives ?|