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2009 Rural Industries Research and Development Corporation. All rights ... 02 6271 4199. Email: [email protected]. ..... Table 1. Plate layout for blanks, standards and sample determination. ...... IREC FARMERS NEWSLETTER No. 171 ...
Rapid Response Screening Tools to Minimise Offsite Impacts of Rice Pesticides RIRDC Publication No. 09/148

RIRDC

Innovation for rural Australia

Rapid Response Screening Tools to Minimise Offsite Impacts of Rice Pesticides

by Anu Kumar, Wendy C. Quayle, Mark Stevens, Marianne Woods and Rai Kookana

September 2009 RIRDC Publication No 09/148 RIRDC Project No. 000810

© 2009 Rural Industries Research and Development Corporation. All rights reserved.

ISBN 1 74151 946 2 ISSN 1440-6845 Rapid Resposen Screening Tools to Minimise Offsite Impact of Rice Pesticides Publication No. 09/148 Project No. 000810 The information contained in this publication is intended for general use to assist public knowledge and discussion and to help improve the development of sustainable regions. You must not rely on any information contained in this publication without taking specialist advice relevant to your particular circumstances. While reasonable care has been taken in preparing this publication to ensure that information is true and correct, the Commonwealth of Australia gives no assurance as to the accuracy of any information in this publication. The Commonwealth of Australia, the Rural Industries Research and Development Corporation (RIRDC), the authors or contributors expressly disclaim, to the maximum extent permitted by law, all responsibility and liability to any person, arising directly or indirectly from any act or omission, or for any consequences of any such act or omission, made in reliance on the contents of this publication, whether or not caused by any negligence on the part of the Commonwealth of Australia, RIRDC, the authors or contributors. The Commonwealth of Australia does not necessarily endorse the views in this publication. This publication is copyright. Apart from any use as permitted under the Copyright Act 1968, all other rights are reserved. However, wide dissemination is encouraged. Requests and inquiries concerning reproduction and rights should be addressed to the RIRDC Publications Manager on phone 02 6271 4165.

Researcher Contact Details Dr Rai Kookana CSIRO Land and Water Waite Road, Urrbrae, SA 5064 Phone: 08 8303 8450 Fax: 08 8303 8565 Email: [email protected] In submitting this report, the researcher has agreed to RIRDC publishing this material in its edited form. RIRDC Contact Details Rural Industries Research and Development Corporation Level 2, 15 National Circuit BARTON ACT 2600 PO Box 4776 KINGSTON ACT 2604 Phone: Fax: Email: Web:

02 6271 4100 02 6271 4199 [email protected]. http://www.rirdc.gov.au

Electronically published by RIRDC in September 2009 Print-on-demand by Union Offset Printing, Canberra at www.rirdc.gov.au or phone 1300 634 313

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Foreword In recent years, increasing awareness of the potential adverse effects of pesticides on the environment has resulted in greater public pressure to assess, monitor and minimise off-site impacts. The application of pesticides for pest control in rice-based cropping systems is commonly practised in Australia. Rapid response tools for quick and cost effective analysis of pesticides needs to be developed to help in quickly identifying problems and intervene in a timely manner should a pesticide be detected in drainage water. Ideally, a rapid response tool with a turn around time of less than one day is needed that can flag the presence of rice pesticides or a marker compound. This is essential for the timely and effective management of the source of the problem. Otherwise, if the analytical results are not readily available, or delayed for more than a week, it is too late to for the irrigation companies or natural resource managers to respond to the problem. The main goal of the project was to develop rapid response tools to allow integrated pesticide management strategies that would lead to the environmental sustainability of rice-farming systems. The importance of this report is that it provides new tools/approaches for analysing pesticides in the environment. Sediment bioassays are usually a relatively simple test that evaluates the response of the tested organism to contaminated sediments under controlled conditions. Through this project, efforts were made to develop a chronic sediment toxicity bioassay that can be used to assess long-term impact of rice pesticides in aquatic ecosystems. Acetylcholinesterase enzyme (AChE) was developed as a biomarker in midges and the sensitivity of pure enzyme from electric eel was also used as rapid and responsive tool for monitoring organophosphate and carbamate pesticides in the environmental samples. Whilst further work is needed to develop AChE enzyme based assay as a more specific method to an individual compound, the available commercial kit can be utilised for non-specific detection of the presence of organophosphate and carbamate pesticides. It is recommended that biomarkers should be used as an integral component of the environmental monitoring programmes where they can be rapidly employed to prioritise tailored pesticide monitoring. Biomarkers should be complemented with chemical analysis, in order to get information on the bioavailable fraction of chemical compounds and to establish a better relation between biomarker responses and chemical data. This project was funded from industry revenue which is matched by funds provided by the Australian Government. This report, an addition to RIRDC’s diverse range of over 1900 research publications, forms part of our Rice R&D program, which aims to improve the profitability and sustainability of the Australian rice industry. Most of RIRDC’s publications are available for viewing, downloading or purchasing online at www.rirdc.gov.au. Purchases can also be made by phoning 1300 634 313.

Peter O’Brien Managing Director Rural Industries Research and Development Corporation

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Acknowledgments The financial support of the RIRDC is gratefully acknowledged. We would like to especially acknowledge Mr Darryl Gibbs and Mr Ian Mason, Chairmen of the RIRDC (Rice) R&D Committee who gave ongoing enthusiastic support for the project. Penny Sloane, Murray Irrigation Ltd, Sigrid Tijss and Rob Kelly, Murrumbidgee Irrigation Ltd and Arun Tiwari, Coleambally Irrigation Co-Op Ltd are thanked for their participation in stakeholder meetings, providing company data and providing cash and in-kind funding support.

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Abbreviations Abbreviation

Meaning

ACh

Acetylcholine

AChE

Acetylcholinesterase

ATC

Acetylthiocholine

CIA

Coleambally Irrigation Co-Op

CV

Coefficient of variation

°C

degrees celsius

DMSO

Dimethyl sulfoxide

DO

Dissolved oxygen

DTNB

5’, 5’-Dithio-bis(2-Nitrobenzoic Acid

EC10

Effect concentration to 10 percent: the concentration of a chemical in water which causes an effect such as malformations in the 10 per cent of the organisms placed in that water for a stated time, usually 48 or 96 h.

EC50

Effect concentration to 50 percent: the concentration of a chemical in water which causes an effect such as malformations in the 50 per cent of the organisms placed in that water for a stated time, usually 48 or 96 h.

EE-AChE

Electric eel-AChE

ELISA

Enzyme Linked Immunosorbent Assay

EPA

Environmental Protection Authority

ESP

Exchangeable sodium percentage

GC/MS

Gas chromatography/Mass spectrometry

g

Grams

h

Hour

IC05

Inhibition concentration at 5 percent: the concentration of a chemical in a sample capable of causing 5% inhibition of activity (such as AChE activity)

IC20

Inhibition concentration at 20 percent: the concentration of a chemical in a sample capable of causing 20% inhibition of activity (such as AChE activity)

Kd

Water adsorption coefficient

L

Litre

μg/L

micrograms per litre

μS/cm

microsiemons per centimetre-conductivity unit

µg/kg

microgram per kilogram

µmol/min/g protein

micromole per minute gram protein- measurement of AChE

mg

Milligram

mg/kg

milligram per kilogram

v

mg/L

milligram per litre

mg/L

milligram per litre

MIA

Murrumbidgee Irrigation Ltd

Min

Minute

mL

Millilitre

mM

milli Moles

mm

Millimetre

NATA

National Association of Testing Authorities

ng/L

Nanogram per litre

NGO

Non-governmental organisation

nM

nano Moles

NOEC

No-Observed Effect Concentration

OP

Organophosphate Pesticide

ppb

Parts per billion

PVC

Polyvinyl chloride

RGA

Ricegrowers’ Association of Australia Inc.

RIRDC

Rural Industries Research and Development Corporation

RT

Room Temperature

SD

Standard Deviation

SE

Standard Error

SEM

Standard Error of the Mean

TSS

Total Suspended Solids

U

Units (arbitrary)

USA

United States of America

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Contents Foreword ................................................................................................................................................ ii Acknowledgments................................................................................................................................. iv Abbreviations......................................................................................................................................... v Contents................................................................................................................................................ vii Tables............................................................................................................................................. viii Figures.............................................................................................................................................. ix Executive Summary............................................................................................................................... x 1. Introduction ....................................................................................................................................... 1 1.1 Objectives ................................................................................................................................... 2 2. Methodology....................................................................................................................................... 3 2.1 Objective 1: Enzyme Linked Immunosorbent Assay (ELISA)................................................... 3 2.2 Objective 2: Suitability of acetylcholinesterase (AChE) enzyme inhibition as a rapid and cost-effective tool....................................................................................................................... 5 2.3 Objective 3: The developmental work of a sediment bioassay based on midge larvae, Chironomus tepperi .......................................................................................................................... 8 3. Results............................................................................................................................................... 11 3.1 Objective 1: Validation of Enzyme Linked Immunosorbent Assay (ELISA) for Environmental Monitoring of Molinate in Agricultural Drainage Water ....................................... 11 3.2 Objective 2: Suitability of acetylcholinesterase (AChE) enzyme inhibition as a rapid and cost-effective tool..................................................................................................................... 17 3.3 Objective 3: The developmental work on sediment bioassay based on midge larvae, Chironomus tepperi ........................................................................................................................ 26 4. Conclusions ...................................................................................................................................... 30 4.1 Rapid analysis using ELISA and Biomarker approach............................................................. 30 4.2 Sediment toxicity assessment.................................................................................................... 30 5. Implications...................................................................................................................................... 31 6. Recommendations............................................................................................................................ 32 7. References ........................................................................................................................................ 33 8. Appendices ....................................................................................................................................... 34

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Tables Table 1. Plate layout for blanks, standards and sample determination. S1, S2 = standard 1, 2 etc. with concentration of molinate (µg/L) below. ......................................................................... 3 Table 2. Relationship between theoretical and measured feeding rates................................................. 9 Table 3. Molinate determined in drainage water using ELISA compared with molinate determined using GC/MS analysis for replicate samples for the year 2005 from a variety of sampling sites in the Murray Valley, NSW. .......................................................................................... 12 Table 4. Normal acetylcholinesterase expression in selected freshwater organisms ........................... 18 Table 5. EC50, EC10 values based on 96 h acetylcholinesterase inhibition in a suite of organisms when exposed to chlorpyrifos ................................................................................................ 18 Table 6. Solvent sensitivity of EE-AChE to three solvents, IC05 with 95% confidence interval in parenthesis.............................................................................................................................. 21 Table 7. Effects of buffer pH on EE-AChE activity over different time intervals............................... 22 Table 8. The 20% inhibition concentration for the 7 pesticides and positive control (eserine) in the EE-AChE assay, 95% CI in parenthesis........................................................................... 23 Table 9. The 20% inhibition concentration for the 7 pesticides and positive control (eserine) for the Abraxis-AChE assay, 95% CI in parenthesis................................................................... 24 Table 10. A series of 48 h acute bioassays with final instar midge larvae under a range of different conditions............................................................................................................................... 28

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Figures Figure 1. Typical molinate ELISA kit standard calibration curve plotted on a standard scale............. 4 Figure 2. Overview of steps for EE-AChE in-vitro assay..................................................................... 7 Figure 3. Overview of steps for Abraxis OP/carbamate-AChE in-vitro assay. .................................... 8 Figure 4. Replicated midge bioassays take place under controlled conditions using field-collected sediment samples................................................................................................................... 9 Figure 5. Typical calibration curves displayed on a logarithmic scale (absorbance at 405 nm) on standards supplied within the commercial molinate ELISA kits, technical molinate standards and Ordram® molinate standards prepared in-house. ......................................... 13 Figure 6. ‘Blank ‘analysis of agricultural drainage waters and ground waters (absorbance at 405 nm) with a range of salinity using commercial molinate ELISA kits........................... 14 Figure 7. Actual concentrations vs. measured concentrations for technical molinate in drainage water samples of varying salinity. ....................................................................................... 15 Figure 8. Actual concentrations vs. measured concentrations for Ordram® molinate in drainage water samples of varying salinity. ....................................................................................... 15 Figure 9. Absorbance (at 405 nm) vs. varying TSS for different concentrations of molinate............ 16 Figure 10. Different levels of AChE inhibition exhibited by midges exposed drainage water samples for 48 h................................................................................................................... 19 Figure 11. EE-AChE activity recorded over 30 minutes at 405 nm of different EE-AChE concentrations...................................................................................................................... 20 Figure 12. Percent inhibition of EE-AChE to eserine (1 mg/L) at 5, 10, 20 and 30 minutes at 405 nm ................................................................................................................................. 20 Figure 13. Percent inhibition of EE-AChE to eserine (0.1 mg/L) at 5, 10, 20 and 30 minutes at 405 nm ................................................................................................................................. 21 Figure 14. Percent inhibition (compared to control) of solvent on EE-AChE activity......................... 22 Figure15. Chemical structures of chlorpyrifos and chlorpyrifos-oxon................................................ 24 Figure 16. AChE inhibition analysis for the river water samples (PR) at varying dilutions (100-12.5%) alone and spiked with chlorpyrifos (CPF) at 1 μg/L (n=2). ........................... 25 Figure 17. AChE inhibition analysis for Milli Q water and river (PR) water samples spiked with chlorpyrifos, diazinon and malathion at their individual IC20 values with a 50% dilution factor (n=2). ........................................................................................................... 25 Figure 18. Emergence success and development times of Chironomus tepperi at 25 ± 1oC under different feeding regimes..................................................................................................... 27 Figure 19. Emergence success of C. tepperi exposed to fenthion-spiked sediment in full life-cycle laboratory bioassays. ........................................................................................................... 28 Figure 20. Development times of C. tepperi exposed to fenthion-spiked sediment in full life-cycle laboratory bioassays. ........................................................................................................... 29

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Executive Summary What the report is about? This report presents the development of rapid response tools for the purposes of integrated pesticide management leading to the environmental sustainability of rice-production systems. This report describes: •

the developmental work conducted for the improvement of an existing immunoassay technique for a commonly used indicator of pesticides (Molinate)



laboratory experiments on the development of cost-effective rapid assessment tools for pesticide detection in drainage water using a biomarker approach (acetylcholinestrase or AChE enzyme)



and chemical and toxicological assessment of sediment associated pesticides in drainage channels.

Who is the report targeted at? This report is particularly relevant to the three irrigation companies supplying water to rice production systems, natural resource managers and the regulatory agencies involved in minimising the adverse impact of rice pesticides. In particular, the project findings are expected to contribute towards maintenance and enhancement of the environmental stewardship of rice production systems with uptake through the Environmental Champions Program of the Rice Industry. Background The rice production system, like other irrigated systems, has to rely on the application of agrochemicals to optimise productivity. This makes it necessary for regulatory authorities to issue water licenses to irrigation companies that enforce and promote careful monitoring, reporting, management, improvement and prevention of pesticide pollution. Currently some of the major constraints of the comprehensive contaminant monitoring programmes that are required to satisfy irrigation water licensing conditions includes the lag-time between environmental sampling and receiving results from analytical laboratories as well as the very high costs of chemical analysis. Historically, molinate herbicide has been used as a useful indicator compound for rice pesticides. Molinate has served the industry very well in reducing the cost of monitoring of pesticides, essentially being used as an early warning system for the presence of other pesticides. However, its manufacture in the USA is being phased out. An alternative rapid response tool with a turn around time of less than one day is needed that can flag the presence of rice pesticides or a marker compound to replace molinate. This is essential for the timely and effective management of the source of the problem. Otherwise, if the analytical results are not readily available, or delayed say for more than a week, then it is too late to for the irrigation companies or natural resource managers to respond to the problem. Aims/objectives The main aim of the project was to develop rapid response tools to allow integrated pesticide management strategies leading to the environmental sustainability of rice-farming systems. The specific objectives were three-fold: 1. Improvement of molinate ELISA (Enzyme Linked Immuno-Sorbent Assays) as a rapid monitoring tool;

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2. To test the suitability of acetylcholinesterase (AChE) enzyme inhibition as a rapid and costeffective tool for monitoring organophosphate and carbamate pesticides in the environmental samples; and 3. The developmental work on sediment bioassay with midge larvae, Chironomus tepperi. Methods used The commercial ELISA kit was evaluated with respect to salinity and turbidity effects on molinate residue analysis, as these are usually the chemical parameters that vary most widely in real drainage water samples. The effect of salinity was evaluated by preparing drainage water samples with a range of salinity (39, 166, 352, 494, 2490, 4210 µS/m) using technical molinate and Ordram molinate. To assess the effects of turbidity on the performance of ELISA kits different quantities of a dispersive soil were added to deionised water and sonicated for 10 minutes to create water with total suspended solids (TSS) of 25, 50, 100, 200, 500 and 1000 mg/L. The evaluation included the analysis of the commercial formulation of molinate i.e. Ordram® in deionised water. Sample analysis was carried out as per the manufacturer’s instructions. Final cell absorbance was measured using either a Molecular Devices EMax plate reader (CSIRO) or a Microplate Manager Biorad Laboratories (Murray Irrigation Ltd) at a wavelength of 450 nM. Corresponding GC/MS analysis was undertaken by a NATA accredited laboratory located in Melbourne, Victoria.

The normal expression of Acetylcholinesterase (AChE) activity in a number of Australian freshwater species such as midges (Chironomus tepperi), shrimp (Paratya australiensis), fish (Melanotaenia fluviatilis) and tadpoles. (Lymnodynastes tasmanieiesis) was established during this study. Midges and shrimp were selected for further in-vivo experiments Acetylcholinesterase inhibition in midges and shrimp were evaluated after 96- h exposure to chlorpyrifos (an organophosphate pesticide) under laboratory conditions. To assess the use of biomarker for environmental samples, AChE activity was determined in midges larvae exposed to the drainage water samples for 48 hours. In-vitro bioassays were developed and validated for measuring AChE inhibition in the environmental samples. Seven compounds were used to validate the in-vitro assays including organophosphate pesticides (chlorpyrifos, diazinon and malathion), carbamate pesticide (carbaryl), thiocarbamate herbicides (molinate and thiobencarb) and a herbicide (clomazone). Laboratory cultures of a midge, Chironomus tepperi were used for ecotoxicological investigations. Acute bioassays (48h) were conducted to compare the toxicity of fenthion (an organophosphate pesticide) to midges under three different conditions. Survival, development and emergence were the endpoints analysed during the fenthion exposures to midge eggs. Results/key findings The commercial molinate ELISA kit was evaluated with respect to salinity and turbidity effects as these are usually the chemical parameters that vary most widely in real drainage water samples. The evaluation included the analysis of the commercial formulation of molinate i.e. Ordram®. The study showed that the determinations of molinate for the commercial product in water may be overestimated using the ELISA calibrations. While we found no significant effect on the response of the ELISA kit by turbidity, our results showed that as the salinity increased, the measured concentrations of molinate herbicide by ELISA also increased which may be higher than the actual values. The significance of this effect may warrant an improved method of calibration of the molinate ELISA method as consistent overestimation can lead to potentially unnecessary subsequent GC/MS analytical costs and reporting. We tested a quick method based on purified AChE enzyme (similar to ELISA) and evaluated it for a range of rice pesticides. We found that the AChE method can detect some pesticides (chlorpyrifos) at

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extremely low (parts per trillion) levels in water within an hour. This is desirable as the environmental acceptance of chlorpyrifos is very low (current notification level of 0.01 µg/L). We also noted that the test responded to thiobencarb and clomazone pesticides if the two compounds were present at elevated levels. We concluded that AChE enzyme based assay can be used as rapid and responsive tool for monitoring rice pesticides in the environmental samples. Whilst further work is needed to develop AChE enzyme based assay as a more specific method to an individual compound, the available commercial kit can be utilised for non-specific detection of the presence of organophosphate and carbamate pesticides. Due to persistent drought and financial constraints, this and other projects were cut short by RIRDC. With the available resources and drought conditions (lack of rice production and the drainage water from rice farms) only the laboratory component of the project could be completed. The two methods evaluated in the project are ready for application under field conditions and demonstration to potential users when the drainage water from rice farms and resources become available. Development of a standardised chronic bioassay for the midge Chironomus tepperi for an integrated assessment of ecotoxicological impacts of pesticides in sediments has been partly completed. However, this would need further development and testing before extending it to the potential users. Implications for relevant stakeholders: 1. ELISA based commercial kits can be utilised more effectively, with proper training on their calibrations, to detect pesticides such as molinate in the rice ecosystems. 2. Biomarker such as AChE inhibition can be used as a diagnostic tool/screening tool ($10/sample) before drainage water samples are sent for pesticide analyses. This will definitely reduce the costs of analyses for the irrigation companies. 3. The growers and irrigation companies have shown keen interest in the area of long-term sustainability and biodiversity. Ecotoxicological assessment of drainage water by using bioassays and a biomarker approach and whole sediment bioassays has the potential to be used as a quick and rapid tool for monitoring rice pesticides by the irrigation companies. Recommendations 1. Tests conducted in this project provide confidence that ELISA and AChE based methods can be used by irrigation companies in-house to screen drainage water samples of key rice pesticides. This approach should be used as a tier one approach and samples showing positive response should be only sent to analytical labs for further confirmation. It is recommended that the use of the above two tools are desirable as screening options. However, there is a need of training of the potential users on their proper calibration to ensure that the results are obtained not only rapidly, but also reliably. 2. The sensitivity of the assay could be enhanced for the some rice pesticides. A passive sampler could be used to concentrate pesticides in the environmental samples. Further trials are needed during the rice growing season to determine if pesticide detection can be improved by preconcentrating samples using passive sampler. 3. Sediment toxicity bioassays have been validated during this project. There is a need to develop more ecotoxicological information on the newer pesticides using native benthic species. This would help in developing water quality and sediment quality guidelines for all rice pesticides. 4. A trial needs to be established in a typical rice growing season to test the suitability of all the techniques developed and validated during this project. These approaches can be used a compliance tool by regulatory authorities.

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5. It is recommended that the uptake of project findings be facilitated through the Environmental Champions Program of Rice Industry.

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1. Introduction All irrigated crops rely on the application of agrochemicals to optimise productivity, and rice based farming systems are no exception. This makes it necessary for regulatory authorities to issue water licenses to irrigation companies that enforce and promote careful monitoring, reporting, management, improvement and prevention of pollution. Currently some of the major constraints of the comprehensive contaminant monitoring programmes that are required to satisfy irrigation water licensing conditions are: 1. The lag-time between sampling and obtaining results from an analytical laboratory; 2. Effective analysis and presentation of large monitoring datasets for ease and transparency of reporting to EPAs, customers and NGOs and for optimisation of water quality management according to irrigation company rice environmental policy; 3. The very high cost of chemical analysis. Across all of the irrigation areas of NSW, contaminant monitoring costs amount to hundreds of thousands of dollars that potentially can be reduced through the development of cheap and rapid monitoring tools and technologies; 4. The limited availability of rapid, off the shelf knowledge and tools for minimization of impacts that may occur from significant contamination events (management and remediation e.g. drainage water pesticide concentrations during high rainfall events, blue-green algal toxicity in farm dams, pathogens and nutrients from irrigated pastures). Currently, molinate is being used as a useful indicator compound for rice pesticides (using ELISA technology). Molinate has served the industry very well in reducing the cost of monitoring of pesticides, essentially being used as an early warning system for the presence of other pesticides. However, its manufacture in the USA is being phased out and may no longer be available for use beyond 2007 to the Australian rice industry. Therefore, there is an urgent need to develop a new indicator tool which enables a rapid response when action levels are exceeded. A previous RIRDC project (UTS-5A), involved analysis of composite daily extracted water samples taken from drainage canals between Griffith and Leeton. These indicated that clomazone reached levels of 6.4 µg/L (Hyne and Aistrop, 2006). Other pesticides such as thiobencarb and atrazine were also detected in drainage waters with diuron, metolachlor and simazine consistently being above ANZECC trigger values for many days in October and November of 2004. Ideally if a rapid response tool (with a turn around time of less than one day) was available that can flag the presence of rice pesticides or a marker compound in drainage water, the source of the problem may be managed much more effectively. Otherwise the results do not become available for more than a week sometimes by which time it is too late to for the irrigation company to respond to the problem. Some pesticides are particularly amenable to detection through rapid response tools. Exposure to pesticides like clomazone, thiobencarb, and chlorpyrifos can inhibit acetylcholinesterase (AChE) activity in invertebrates and fish. Preliminary studies from the RIRDC project (CSL-16A) have shown that exposure to rice drainage water can cause AChE activity inhibition in midges. During a workshop with stakeholders, it was agreed that AChE, which is specific to certain pesticides used in rice production, can potentially be developed as a very useful rapid response tool for rice pesticides. Similarly commonly detected pesticides such as diuron, and triazines (non rice pesticides), respond to rapid field tests based on ELISA technology. These tests are available but need to be optimised. These single-day response tests of drainage water can flag if one or more of these pesticides are present in the drainage water. Only water samples with positives during the first screen need to go for pesticide

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analyses to confirm the pesticide residues. This way, the industry can cut down on the number of samples selected for chemical monitoring in the drainage water, making it very cost effective. The rapid monitoring will therefore facilitate better management of pesticide issues as the new tools will allow an immediate intervention in situations where a pesticide has migrated off-site. Timeliness of intervention is the key factor in better management or minimising impact on environment.

1.1 Objectives The key objective of the study was to develop rapid response tools that facilitate integrated pesticide management strategies to further the environmental sustainability of rice-farming systems. Originally the specific objectives of the project were: •

Developing cost-effective, rapid assessment tools for pesticide detection in the drainage water using a biomarker approach.



Conducting chemical and toxicological assessment of sediment associated pesticides in the drainage channels.



Enhancing the "clean and green" image of rice production systems through linkages with the Environmental Champions Program.

The funding to the project stopped prematurely in June 2008, due to the persistent drought conditions severely restricting the productivity of the Australian rice industry since 2004. Due to the curtailing of the project duration, the above objectives had to be modified in consultation with RIRDC and other stakeholders. These specific activities were undertaken to optimise the available resources: 1. Improvement of molinate ELISA (Enzyme Linked Immuno-Sorbent Assays) as a rapid monitoring tool; 2. To test the suitability of acetylcholinesterase (AChE) enzyme inhibition as a rapid and costeffective tool for monitoring organophosphate and carbamate pesticides in the environmental samples; and 3. The developmental work on sediment bioassay based on midge larvae, Chironomus tepperi exposed to an organophosphate pesticide. The report is organised to discuss each of the above specific activity/objective one by one under the methodology and results sections followed by conclusions, implications and recommendations arising from the project.

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2. Methodology 2.1 Objective 1: Enzyme Linked Immunosorbent Assay (ELISA) Commercially available ELISA kits for molinate analysis (Abraxis) were evaluated. Sample analysis was carried out as per the manufacturer’s instructions (Appendix 1). Final cell absorbance was measured using either a Molecular Devices EMax plate reader (CSIRO) or a Microplate Manager Biorad Laboratories (Murray Irrigation Ltd) at a wavelength of 450 nm with the plate layout shown in Table 1. Calibration curves were developed using standards all ready prepared in the kits (Figure 1) or by making corresponding standards using technical molinate (Dr Ehrenstorfer, Germany, 99% purity) or commercial molinate formulation Ordram® in deionised water or authentic drainage water samples and groundwater by standard methods. Corresponding GC/MS analysis was undertaken by a NATA accredited laboratory located in Melbourne, Victoria. Table 1. Plate layout for blanks, standards and sample determination. S1, S2 = standard 1, 2 etc. with concentration of molinate (µg/L) below. X1, X2 etc = unknowns 1

2

3

4

5

6

7

8

9

10

11

12

A

-

-

S1 0.000

S1 0.000

S2 1.25

S2 1.25

S3 2.500

S3 2.500

S4 10.00

S4 10.00

S5 25.00

S5 25.00

B

S6 100.0

S6 100.0

X1 1.000

X1 1.000

X2 1.000

X2 1.000

C D E F G H

2.1.1 Collection and preparation of samples to assess the effects of turbidity on the performance of ELISA kits A soil sample was collected from the Wakool region of the Western Murray Valley in an area known to have dispersive sodic soils (ESP> 15%). The soil belongs to the Great Soil Group Grey and Brown Soils with heavy texture (up to 80% clay). The sample collected was from the bank of an irrigation channel. The sample was air dried and different quantities of soil were added to deionised water and sonicated for 10 minutes to create water with total suspended solids (TSS) of 25mg/L, 50 mg/L, 100 mg/L, 200 mg/L, 500 mg/L and 1000 mg/L. The top molinate standard in a commercial ELISA kit was used to spike each TSS sample to concentrations of 1.25 µg/L, 2.5 µg/L, 10 µg/Land 25 µg/L of molinate. A blank TSS solution was analysed for the range of TSS outlined above containing no molinate standard. A further calibration containing no soil and using the 0 – 25 µg/L ELISA standard range spiked into deionised water was also constructed.

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Figure 1. Typical molinate ELISA kit standard calibration curve plotted on a standard scale.

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2.2 Objective 2: Suitability of acetylcholinesterase (AChE) enzyme inhibition as a rapid and cost-effective tool 2.2.1 In-vivo exposures: AChE Inhibition in a suite of aquatic organisms 2.2.1.1 Choice of the pesticide Chlorpyrifos was selected for carrying out in-vivo pesticide exposure studies. This was an appropriate choice, as it is an organophosphate pesticide with AChE inhibition as its mode of action, is commonly used in rice production system and being very toxic its acceptance in the environment is very low. Therefore a sensitive assay is desirable for this pesticide. 2.2.1.2 Pesticide exposure testing The toxicity tests consisted of eight treatments including a control and a solvent control. Bioassays were conducted using shrimp, Paratya australiensis (10–15 mm long), tadpoles of grass-spotted frog, (Lymnodynastes tasmanieiesis) (96 h old), Murray rainbowfish, Melanotaenia fluviatilis (2 months old) and midge 4th instar larvae, Chironomus tepperi (approximately 4 days after hatching). The tadpoles, shrimp and fish bioassays were conducted for 96 h where as midges were exposed to chlorpyrifos for 48 h. The pesticide concentrations used in the definitive experiments were based on range-finding tests. During range-finding tests, pesticide concentrations leading to 100% mortality of test organisms after 96 h pesticide exposures were not included for AChE inhibition experiments. The tests were conducted using four replicates of each treatment. For shrimp and fish bioassays, each replicate consisted of a 1 L beaker that contained 800 mL of test solution and ten randomly allocated shrimp or fish fry. Midge and tadpole bioassays were conducted in 25 mL of test solutions. The test containers were covered with plastic film to reduce volatilisation and then randomly allocated in an incubator set at 23 ± 1oC with a light intensity of approximately 800 Lux at the surface of the water and a 16:8 h light:dark regime. Daily measurements were made of pH, electrical conductivity and dissolved oxygen. The toxicity tests were considered valid if the mortality in each of the diluent water controls and solvent controls was not greater than 20% and the dissolved oxygen (DO) concentration in all the test solutions remained greater than 60% saturation. After 48 and 96 h exposures, the test organisms were frozen at -70ºC for further AChE analyses. 2.2.1.3 AChE analyses The interpretation of AChE activity depends on the knowledge of “normal” levels. The normal expression of AChE activity in a number of Australian freshwater species such as midges, shrimp, fish and tadpoles was measured as a first step. After 96 h exposures to chlorpyrifos, the surviving tadpoles, midges, fish and shrimp in different concentrations of pesticides and control treatments were frozen at –70°C for AChE analyses. For AChE analyses, each individual shrimp and fish were homogenised for AChE analyses. Since the masses of both midges and tadpoles are small, 2-3 individuals were pooled together for AChE analyses. Tissue samples were prepared in 1.5 mL Eppendorf tubes and homogenized in 0.3 mL phosphate buffer (pH 8). The tissue samples were placed on ice immediately to maintain enzyme stability and were used for both AChE activity measurements and protein analysis. AChE activity was measured spectrophotometrically (405 nm) by following the increase of the yellow anion, 5-thio-2nitrobenzoate, which was produced when thiocholine reacts with dinitrobenzoic acid (DTNB). This change in colour was measured using a scaled-down version of the method described by Ellman et al. (1961), using a Titertek® Multiskan MC Photometer, which allowed large numbers of samples to be analysed at one time. Protein content in tissues was determined as described by Lowry et al. (1951).

5

AChE activity was standardised to protein concentration in each sample. AChE activity was expressed as micromoles per minute per gram protein (μmoles/min/g protein). AChE inhibition was calculated as a percentage of control values.

2.2.2 Drainage water toxicity: AChE Inhibition in midges AChE biomarker can be used as a biomarker for evaluating the toxicity of the drainage water samples. For this approach, laboratory cultured organisms maintained in controlled conditions were required. Briefly, around 110 drainage water samples were collected from the rice growing regions. These water samples were collected by the irrigation companies in 2004-2005 and were transported to the Aquatic Ecotoxicology laboratory in Adelaide. Laboratory cultured midge larvae were exposed to the drainage water for 48 h. Three to four replicates were set-up for each drainage water samples. After 48 h exposures, these midges were frozen at –70°C for AChE analyses.

2.2.3 In-vitro AChE assay - Sensitivity to seven pesticides The assay is a qualitative, colorimetric assay (modification of the Ellman method) for the detection of organophosphates (OP) and carbamates using a purified enzyme (AChE). It is based on the ability of OPs and carbamates to inhibit AChE. AChE hydrolyses acetylthiocholine (ATC) to thiocholine, which reacts with 5’,5’-Dithio-bis(2-Nitrobenzoic Acid) (DTNB) to produce a yellow colour which is read at 405 nm using a spectrophotometer. A sample containing an OP or carbamate will inhibit AChE reducing or eliminating colour formation. A positive result is indicated at 20% inhibition (IC20) of colour development. Two different AChE assays were conducted, one with the electric eel (Electrophorus electricus) (EEAChE) and the other a commercially available kit, Abraxis OP/carbamate assay with AChE purified from insect cells. Trials were conducted to determine: 1. Optimal EE-AChE concentration, 2. Solvent sensitivity of EE-AChE, 3. Effects of buffer pH (pH 8.0 vs pH 7.0) in EE-AChE assay, 4. Sensitivity of EE-AChE to 7 pesticides (with and without oxidation for OPs), 5. Sensitivity of Abraxis kit-AChE to 7 pesticides (with oxidation), and 6. River water sample All solvents used for stock solutions were analytical grade. Acetylcholinesterase from electric eel (Electrophorus electricus) (EE-AChE) type V-S, > 1000 units/mg protein, albumin from bovine serum (BSA), acetylthiocholine iodide (ATC), 5’,5’-Dithio-bis(2-Nitrobenzoic Acid) (DTNB), carbaryl (99.8% purity), chlorpyrifos (99.5% purity), clomazone (97.0% purity), diazinon (99.0% purity), eserine, malathion, molinate (99.0% purity), thiobencarb (99.8% purity) were purchased from SigmaAldrich Pty. Ltd. The OP/C screen kit was purchased from Abraxis, LLC. 2.2.3.1 Stock solutions The assay was conducted with phosphate buffer (0.1 M, pH 8.0). Stock solutions of ATC (2 mM) and DTNB (2.4 nM) were prepared fresh daily in phosphate buffer. Stock solutions of EE-AChE were prepared in 1 mg/mL BSA and stored at 4°C. Dilute stock solutions of EE-AChE (< 1 mg/mL) were prepared weekly in 1 mg/mL BSA. Pesticide stock solutions were prepared in acetone (4-5 g/L) and

6

stored at -18°C. Working stock solutions were prepared weekly in 50% methanol. All solutions in the Abraxis OP/C kit were prepared to the manufacturer’s instructions. 2.2.3.2 EE-AChE assay All solutions were allowed to come to room temperature before commencement. Samples were run in duplicate, together with positive controls (eserine) and a negative control (50% methanol). The detection limits for OP and carbamates varied depending on their ability to inhibit AChE. Figure 2 is an overview of the steps involved in the assay. The commercially available OP/carbamate 96 well Abraxis kit was used to compare the pesticide sensitivities obtained with the EE-AChE. This kit uses AChE obtained from insect cells (unknown species) and an overview of the methodology is outlined in Figure 3.

Assay Buffer

Standards / Sample

AChE

15 min @ RT Substrate (ATC)

Chromogen (DTNB)

Read at 405 nm

30 min @ RT

Figure 2. Overview of steps for EE-AChE in-vitro assay. RT (Room temperature)

7

Assay Buffer Standards / Sample Oxidiser 5 min @ RT Neutraliser AChE 15 min @ RT Substrate (ATC) Chromogen (DTNB) 30 min @ RT Stopping Solution Read at 405 nm Figure 3. Overview of steps for Abraxis OP/carbamate-AChE in-vitro assay. RT (room temperature)

2.3 Objective 3: The developmental work of a sediment bioassay based on midge larvae, Chironomus tepperi 2.3.1 Laboratory midge cultures Laboratory cultures of Chironomus tepperi Skuse (Diptera: Chironomidae) were established using the techniques of Stevens. Cultures were maintained at 25±1oC with a 15:9 light:dark photoperiod.

2.3.2 Containers, substrate and solution Bioassays were conducted using 600 mL capacity glass beakers (internal diameter 85 mm, Crown Boroglass, Crown Scientific, Sydney). Twenty five beakers were used in total (5 replicates of 5 feeding rates). To determine the optimal supplemental feeding rate it was essential that the substrate contained no endogenous food sources, so a sand substrate was used. River sand was washed for 20 minutes using tap water with constant agitation, oven dried, and passed through a 710 µm sieve to remove larger grains. The sand was then oven dried, prior to being autoclaved at 121oC for 20 minutes, oven dried again, and allowed to cool. One hundred grams of sand was then placed in each beaker. Martin’s 1x rearing solution was then added to give a total volume (sand plus rearing solution) of 400 mL.

8

2.3.3 Food and feeding rates A food suspension was prepared by hand-grinding K9® brand goldfish food (Carnation, Melbourne, 8 to 9% moisture) as finely as possible and adding 2 g to 125 mL of deionised water in a 250 mL laboratory bottle (Schott). The 16 mg/mL suspension was shaken for 5 minutes and stored at 4oC until use. The five feeding rates chosen for evaluation ranged from 0.075 to 0.6 mg food/larvae/day, however rapid settling of larger particles from the food suspension suggested that smaller amounts may have actually been added. To evaluate this, aliquots of food were pipetted into pre-weighed glass vials, dried overnight at 105oC, and reweighed, allowing food content to be calculated on a dry weight basis. Results of this study are shown in Table 2.

2.3.4 Experimental conditions, test insects, duration and monitoring The beakers were randomly allocated to the 5 different feeding rates, labelled, and randomised across 4 shelves in a controlled temperature room set at 25oC (±1.5). Lighting was provided by a mixture of incandescent and cool fluorescent globes positioned opposite the shelving and set to provide a 15:9 light:dark cycle. A single C. tepperi egg mass showing advanced egg development was selected from a laboratory culture maintained at Yanco Agricultural Institute, and 15 eggs were dissected from the egg mass and placed in each beaker. The first aliquot of food was added to the beakers and covered with plastic film. Aeration was provided to each beaker via a 200 µL plastic pipette tip attached via thin plastic tubing to a circular air line connected to two aquarium aerators (Figure 4). Table 2. Relationship between theoretical and measured feeding rates. Volume of suspension (mL)*

Theoretical food content (mg)

Theoretical feeding rate (mg/larva/day)

Measured feeding rate (mg/larva/day)

1.125

18.0

0.60

0.48 ± 10%

0.844

13.5

0.45

0.33 ± 14%

0.562

9.00

0.30

0.22 ± 14%

0.282

4.50

0.15

0.13 ± 13%

0.141

2.25

0.075

0.06 ± 25%

*16 mg/mL, 48 h feeding intervals

Figure 4. Replicated midge bioassays take place under controlled conditions using fieldcollected sediment samples.

9

Additional food aliquots were added every 48 h, with prepared food being discarded and replaced after every second feeding to minimise the risk of settling and declining suspension volume leading to progressive increases in actual food quantities delivered. Emergence (including the sex of emerged adults) was recorded twice daily, with the time to emergence being considered as the time between initiation of the experiment and the midpoint between the two observations between which emergence occurred. Emergence was considered to have occurred if the midge had completely separated from the pupal exuvium. At the conclusion of the experiment the contents of each beaker was tipped into a white plastic tray and checked for any surviving larvae or pupae.

2.3.5 Validation of bioassay using fenthion The vast majority of toxicity tests are conducted using highly purified laboratory water under highly controlled conditions. The objective of this study was therefore to determine the effects of spiked sediments on the toxicity and bioavailability of fenthion to the midges. Information on environmental fate and physico-chemical properties of fenthion can be accessed at the following web-site: http://extoxnet.orst.edu/pips/fenthion.htm. Acute bioassays (48 h) were conducted to compare the toxicity of fenthion to midges under three different conditions. In the first treatment, 4th instar midge larvae were exposed to a series of fenthion concentrations in clean water with no sediments. In the second treatment, midges were exposed to fenthion via spiking sediments to a series of fenthion concentrations. In the third treatment, clean sediments were added to each test beaker and then the overlying water was spiked to a range of fenthion concentrations during 48 h bioassays. Forty eight hour LC50 values were compared between different treatments. The response of chironomid larvae to pesticides depends on their age at pesticide exposure, with larvae at earlier developmental stages being more susceptible. Testing across a complete life cycle (from egg to adult emergence) can provide a more sensitive test than relying on short duration 4th instar bioassays. To test this, clean sediment was spiked with fenthion at different concentrations, C .tepperi eggs were added, and their survival and development was monitored. This was done under the optimised feeding conditions developed during the experiment described previously.

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3. Results 3.1 Objective 1: Validation of Enzyme Linked Immunosorbent Assay (ELISA) for Environmental Monitoring of Molinate in Agricultural Drainage Water 3.1.1 Performance of Molinate ELISA kits using kit standards The performance of the molinate ELISA kits used as per manufacturers instructions in an irrigation company laboratory were assessed. Murray Irrigation Ltd. tested drainage water samples weekly from the 28/9/06-13/12/06. The tests were precise based on the reproducibility of duplicates of the standard working range (1.25 – 100 µg/L). Within any individual kit the mean %CV was typically ethanol>methanol (Table 6; Figure 14). As methanol exhibited the least toxicity to the EE-AChE it was used as the diluent for the pesticide trials and was used as the negative control. Table 6. Solvent sensitivity of EE-AChE to three solvents, IC05 with 95% confidence interval in parenthesis. Solvent

IC05 (95% CI)

Methanol

45.6 (40.7-51.0) %

Ethanol

28.6 (14.5-56.5)%

DMSO

2.40 (0.52-11.1) %

21

Methanol Ethanol DMSO

100

% Inhibition

80

60

40

20

0 1.9

9.5

19.0

47.5

100.0

% Solvent

Figure 14. Percent inhibition (compared to control) of solvent on EE-AChE activity. Bar represents the mean of 3 individual trials, error bars represent the standard error of the mean (n=9).

3.2.2.3 Effects of buffer pH (pH 8.0 v’s pH 7.0) in EE-AChE assay The effect of buffer pH on sensitivity of the EE-AChE was considered. Although the sensitivity was enhanced in assays using buffer pH 7.0 (approximately 10% more inhibition), this was also observed with the solvent controls. Therefore, when the solvent effects (negative control) were accounted for the differences in sensitivities were negligible (Table 7). Table 7. Effects of buffer pH on EE-AChE activity over different time intervals

Percent inhibition at different time intervals (min)

Eserine 1 mg/L Eserine 0.1 mg/L

5

10

15

20

25

30

pH 7.0

61

65

67

68

70

71

pH 8.0

62

66

67

69

69

70

pH 7.0

22

23

24

23

22

21

pH 8.0

18

20

21

20

19

18

3.2.2.4. Sensitivity of EE-AChE to 7 pesticides (with and without oxidation for OPs) Seven compounds were used to validate the EE-AChE in-vitro assay including organophosphate pesticides (chlorpyrifos, diazinon and malathion), carbamate pesticide (carbaryl), thiocarbamate herbicides (molinate and thiobencarb) and a herbicide (clomazone). Initially, range finding tests were conducted over several concentration ranges and definitive tests were conducted once the effective range was obtained. The 20% inhibition concentration (IC20) was calculated for each compound (where possible). The OPs were also tested with an oxidation step to convert the parent compound to their active oxon-analogues.

22

The EE-AChE in-vitro assay could only detect the test compounds in the mg/L range with the EEAChE displaying the highest sensitivity to carbaryl (1.5 mg/L) (Table 8). With oxidation, the sensitivity of EE-AChE to the OPs increased significantly. This was especially evident for chlorpyrifos where the IC20 decreased nearly 430 times. Table 8. The 20% inhibition concentration for the 7 pesticides and positive control (eserine) in the EE-AChE assay, 95% CI in parenthesis.

Compound

IC20 (mg/L)

IC20 (with oxidation) (mg/L)

Eserine

0.153 (0.134-0.175)

Not analysed

Chlorpyrifos

53.6 (49.2-58.3)

0.125 (116.0-134.8)

Diazinon

> 500

4.5 (3.8-5.3)

Malathion

> 10

0.227 (201.6-256.4)

Carbaryl

1.5 (1.2-1.8)

Not analysed

Molinate

388.7(284.8-530.4)

Not analysed

Thiobencarb

> 500

Not analysed

Clomazone

> 500

Not analysed

3.2.2.5. Sensitivity of Abraxis-AChE to 7 pesticides (with oxidation) The same seven compounds were used to validate the Abraxis-AChE in-vitro assay as the EE-AChE assay. The 20% inhibition concentration (IC20) was calculated for each compound. For all compounds, oxidation and neutralisation steps were conducted. The Abraxis-AChE was highly sensitive in the detection of the OPs. The IC20 concentrations were in the μg/L range with the Abraxis-AChE displaying the highest sensitivity to chlorpyrifos (0.5 μg/L) (Table 9). When compared to the EE-AChE assay, the enzyme in the Abraxis-AChE assay is significantly more sensitive toward the detection of OPs and carbamates. This is especially evident for the phosphorothionates (those compounds that require oxidation to become potent AChE inhibitors). For example, for diazinon the EE-AChE assay failed to detect a difference in inhibition at concentrations up to and including 500 mg/L, where as the Abraxis-AChE assay could detect this compound at levels as low as 1.5 μg/L (≥ 830,000 x more sensitive). The difference in sensitivities of the carbamate pesticides, thiocarbamate herbicides and herbicides between the two assays was not as significant although the Abraxis-AChE assay was more sensitive.

23

Table 9. The 20% inhibition concentration for the 7 pesticides and positive control (eserine) for the Abraxis-AChE assay, 95% CI in parenthesis

Compound

IC20 (μg/L)

Eserine

357.7 (231.1-553.6)

Chlorpyrifos

0.51 (0.38-0.69)

Diazinon

0.64 (0.51-0.80)

Malathion

1.5 (0.76-3.1)

Carbaryl

233.6 (194.3-281.0)

Molinate

20,400 (9,500-43,800)

Thiobencarb

110,600 (85,700-142,800)

Clomazone

202,800 (160,900-255,500)

The sensitivity of the assay could be enhanced for the OPs with the addition of an oxidiser. The phosphorothionates OPs are lipophilic with one thione moiety (P=S) and three –OR groups attached to phosphorus atom (Figure 15). The oxidised analogues are more polar characterised with a double phosphorus oxygen bond (P=O). Generally, the oxon-analogues are more potent AChE inhibitors than the parent compound (Kralj et al. 2006). The oxidiser used in the Abraxis kit is unknown. A study by Kralj (2006) reported the use of N-bromosuccinimide (NBS), hypochloride, oxone, hydrogen peroxide and others for the oxidation of thio-OPs in aqueous solutions. Further trials are needed to determine the suitability of some of these compounds as oxidisers for the use in the current assay to improve the sensitivity.

Figure15. Chemical structures of chlorpyrifos and chlorpyrifos-oxon.

3.2.2.6 River water sample Tests were conducted with a river water sample with an oxidation step and without the oxidation step. No significant differences were noted for samples that were oxidised and those that were not, except in the chlorpyrifos spiked samples (p=0.05). In these samples, no AChE inhibition was noted indicating the need to convert chlorpyrifos to its active oxon analogue for AChE inhibition to occur. No AChE inhibition was noted for the river water samples. Those samples spiked with 1 μg/L chlorpyrifos exhibited a significant increase in AChE inhibition However, there was no significant difference between the spiked Milli Q water sample and the river water sample at the concentrations tested (p=0.05). This indicates that there were no matrix interferences for detecting chlorpyrifos in this water sample (Figure 16).

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Alone 100

CPF spike with 1 ug/L

90 80

% Inhibition

70 60 50 40 30 20 10 0 Milli Q water

PR 100%

PR 50%

PR 25%

PR 12.5%

Concentration

Figure 16. AChE inhibition analysis for the river water samples (PR) at varying dilutions (10012.5%) alone and spiked with chlorpyrifos (CPF) at 1 μg/L (n=2).

Assays were conducted with spiked river water samples spiked with chlorpyrifos (0.5 μg/L), diazinon (0.64 μg/L) and malathion (1.5 μg/L) at their individual IC20 values. This was compared to a Milli Q water samples spiked with the same compounds. No significant difference was noted between the river water extracts and the Milli Q water, indicating that no matrix effects were present in the water samples (Figure 17). Milli Q water 100

PR water

90 80

% Inhibition

70 60 50 40 30 20 10 0 IC20

IC10

IC5

IC2.5

IC1.25

Concentration

Figure 17. AChE inhibition analysis for Milli Q water and river (PR) water samples spiked with chlorpyrifos, diazinon and malathion at their individual IC20 values with a 50% dilution factor (n=2).

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3.2.3 Summary Based on the preliminary results, it can be included that specific biomarkers of exposure, such as acetylcholinesterase (AChE) inhibition used in this study, can be used as rapid and responsive tool for the assessment of toxic effects caused by selected rice pesticides. The knowledge of sources of variability in AChE measurement is an important factor to be taken into account for the optimisation and standardisation of protocols. Factors such as sampling methods and storage, preparation of reactive agents, preparation, processing and storage of tissues, preparation of reactive agents, physicochemical conditions of measurement and instrumentation needs to be taken into consideration for routine application of this biomarker.

3.3 Objective 3: The developmental work on sediment bioassay based on midge larvae, Chironomus tepperi 3.3.1 Feeding rate Results of the feeding rate study are summarised in Figure 18. Progressively increasing the feeding rate from 0.141 to 1.125 mL per 48 h did not significantly affect emergence success (Figure 18 a), which averaged 63% across all treatments. Development times were significantly affected for both males and females (Figure 18 b, c) and decreased with the provision of additional food. Significant (p < 0.05 decreases in development times for males did not occur at rates above 0.844 mL / 48 h, whilst significant decreases for females did not occur at feeding rates above 0.566 mL / 48 h. A nonsignificant downward trend after 0.566 mL/48 h continued up to the highest feeding rate evaluated. Results of this study show that the optimal feeding rate for 15 C .tepperi larvae in our bioassay system is in excess of 0.566 mL / 48 h, but that rates above 0.844 mL / 48 h provide only small and nonsignificant increases in development rates. There was no evidence of accumulation of excess food at the 0.844 mL / 48 h feeding rate until the majority of midge emergence had occurred, however uneaten food started to accumulate at the 1.125 mL / 48 h rate from relatively early during the bioassays. On the basis of this result a feeding rate of 0.9 mL / 48 h was chosen as the uniform feeding rate for use in all subsequent bioassays.

26

100

(a) emergence success (both sexes)

90

nsd

% emergence + SE

80 70 60 50 40 30 20 10 0 0.141

0.282

0.566

0.844

1.125

development time (hrs)+ SE

feeding rate (mL suspension / 48 hrs) 600 500

(b) development time (replicate means) males

a

400

b

300

b c

200

c

100 0 0.141

0.282

0.566

0.844

1.125

development time (hrs) + SE

feeding rate (mL suspension / 48 hrs) 600

(c) development time (replicate means) females

a

500 400

b c

300

c c

200 100 0 0.141

0.282

0.566

0.844

1.125

feeding rate (mL suspension / 48 hrs)

Figure 18. Emergence success and development times of Chironomus tepperi at 25 ± 1oC under different feeding regimes. nsd, no significant difference between treatments (P > 0.05). Within each graph columns with different letters are significantly different (ANOVA, LSD test, P < 0.05).

3.3.2 Validation of Midge bioassays: toxicity to Fenthion To validate the technique, we conducted a series of replicated experiments looking at the response of midge larvae to sediments spiked with the organophosphate insecticide fenthion. Significant suppression of emergence success occurred at fenthion concentrations of 600 µg/kg and above. The effect of fenthion concentration on the development times of midges that successfully developed to adults was less dramatic, although the overall trend was for delayed development at higher concentrations. This result may reflect low persistence of fenthion in the bioassay system – either individuals died shortly after exposure began, or survived the initial exposure and were minimally affected thereafter. As was anticipated, the highest level of acute toxicity occurred when the chemical was delivered in water without a soil substrate being present. When the water was spiked above clean soil, toxicity declined markedly, and the lowest toxicity was observed when the soil itself was spiked and overlayed

27

with clean water (Table 10). This last situation is analogous to the system used in the chronic bioassays, and the important point to note here is that the chronic full life-cycle bioassay demonstrated a significant effect at half the soil fenthion concentration required to obtain 50% mortality in the corresponding acute assay with final instar larvae (Figure 19), demonstrating the improved sensitivity of the chronic assay. Development times of C. tepperi exposed to fenthion-spiked sediment in full lifecycle laboratory bioassays was variable among male and female midges (Figure 20). Further validation work may show greater improvements in sensitivity with compounds (such as some herbicides) that do not have strong acute effects at low concentrations. Table 10. A series of 48 h acute bioassays with final instar midge larvae under a range of different conditions LC50 (ppb)

LC90 (ppb)

No soil present

163

319

Spiked water*

484

1200

Spiked soil**

1174

2313

Conditions

Note: *1:4 soil:water ratio spiked into surface water post-flood, **1:4 soil:water ratio, 6 reps x 5 rates, n = 60 at each rate ppb=parts per billion; µg/kg in soil or µg/L in water

120

emergence success (%)

100

a a

a a

a

80

b

60

b

40

b 20

0 control

200

300

400

500

600

700

800

sediment fenthion concentration (µg/kg)

Figure 19. Emergence success of C. tepperi exposed to fenthion-spiked sediment in full lifecycle laboratory bioassays. Columns with different letters are significantly different (p < 0.05).

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340

male developmnent time (hrs)

320 300 280 260 240 220 200 180 160 140 control

200

300

400

500

600

700

800

sediment fenthion concentration (µg/kg) 340

female developmnent time (hrs)

320 300 280 260 240 220 200 180 160 control

200

300

400

500

600

700

800

sediment fenthion concentration (µg/kg)

Figure 20. Development times of C. tepperi exposed to fenthion-spiked sediment in full lifecycle laboratory bioassays.

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4. Conclusions 4.1 Rapid analysis using ELISA and Biomarker approach Overall, the results from the investigations support the view that for the routine monitoring of molinate in agricultural drainage water quality ELISA method can be used. The variations seen in the natural samples compared with kit standards do not merit the application of a correction factor for further optimisation. However, training of the operator in proper calibration of ELISA method is desirable. Indeed, owing to the rapid analysis of samples after collection using the ELISA assays and with very little transport or storage involved, it is considered here that ELISA results may be more representative of field concentrations than those analysed by GC/MS techniques in laboratories remote from field sampling sites. Based on the preliminary results, it can be concluded that specific biomarkers of exposure, such as acetylcholinesterase (AChE) inhibition used in this study, can be used as rapid and responsive tool for the assessment of toxic effects caused by selected rice pesticides. Whole-body AChE activity measurement in C. tepperi 4th instar proved to be a sensitive bioindicator for the presence of OP and carbamate pesticides in the drainage water. However, this would require the establishment of ecotoxicological facilities at the irrigation companies for maintaining cultures of midges under controlled conditions. An alternative approach is to use the commercially available kit as a simple, robust and reliable biomarker in bio-monitoring programmes for OP and carbamate pesticides in the rice ecosystems AChE biomarker can be used by irrigation companies for screening drainage water samples for the presence of organophophate and carbamate pesticides. This use of biomarkers approach can facilitate a timely intervention in terms of management of pesticide residues, should a pesticide is detected in drainage water. Samples that show positive response could then be sent to commercial analytical laboratory for confirmation of the presence of pesticides in the environmental samples. This tiered approach would help the irrigation companies and the regulators to manage pesticide detection issues in the agricultural systems in a more efficient manner.

4.2 Sediment toxicity assessment Studying the response of the biota to contaminants can complement chemical analyses and significantly enhance the predictive capabilities of monitoring programs. While chemical analyses represent an important component of monitoring programs, sole reliance on this approach may not effectively predict the impact of pesticides on a receiving system. The key to whether a contaminant will affect an organism is bioavailability, or the extent to which the contaminant can enter the organism. In reality, complex mixtures of pesticides can result in a combined toxicity very different from that predicted for the individual components alone. No analytical method at present can effectively predict this interaction and the associated toxic effects. However, biological data may greatly augment chemical data by indicating the availability of contaminants and the potential for negative effects on receiving systems. The midge bioassay developed during this project has the potential to be used as a standard ecotoxicological test for monitoring adverse impacts due to pesticide use in agricultural systems. Chironomus tepperi is a sensitive species and can be routinely used in pesticide monitoring programs.

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5. Implications 1. ELISA based commercial kits can be utilised more effectively, with proper training on their calibrations, to detect pesticides such as molinate in the rice ecosystems. 2. Specific biomarkers of exposure, such as AChE inhibition used in this study, proved to be responsive tools for the assessment of toxic effects caused by the OP and carbamate pesticides present in the drainage water samples. Biomarkers such as AChE inhibition has potential for being incorporated into a regional monitoring program. It can be used as a diagnostic tool/screening tool before drainage water samples are sent for pesticide analyses. This will definitely reduce the costs of analyses for the irrigation companies. AChE will respond to rice pesticides and can be used as a biomarker for $10 per sample by using a commercial kit. This biomarker can also be used by irrigation companies to test relationships between changing patterns of pesticide application. In-vivo and in-vitro methods trialled and validated in this study can provide guidance on the safe discharge of drainage water into the receiving environment. The AChE biomarker can also be used for assessing in stream toxicity and for run-off mitigation programs. 3. The growers and irrigation companies have shown keen interest in the area of long-term sustainability and biodiversity. Ecotoxicological assessment of drainage water by using bioassays and a biomarker approach and whole sediment bioassays has the potential to be used as a quick and rapid tool for monitoring rice pesticides by the irrigation companies.

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6. Recommendations 1. Tests conducted in this project provide confidence that ELISA based method can be used by irrigation companies in-house to screen drainage water samples of key rice pesticides. This approach should be used as the first stage of a tiered approach and samples showing positive responses should only be sent to analytical labs for further confirmation. 2. The knowledge of sources of variability in AChE measurement is an important factor to be taken into account for the optimisation and standardisation of protocols. Factors such as sampling methods and storage, preparation of reactive agents, preparation, processing and storage of tissues, preparation of reactive agents, physico-chemical conditions of measurement and instrumentation needs to be taken into consideration for routine application of this biomarker. Standard operating protocols can be developed for the users and tailored workshops can be organised for the irrigation companies to address these issues. 3. The sensitivity of the assay could be enhanced for the some rice pesticides. For example, a passive sampler could be used to concentrate pesticides in the environmental samples. Further trials are needed during the rice growing season to determine if pesticide detection can be improved by sample pre-concentrating. 4. Sediment toxicity bioassays have been validated during this project. There is a need to develop further ecotoxicological data on a suite of organisms for the rice pesticides, particularly for newly developed pesticides and in sediments. Ecotoxicological assessment using native benthic species would also provide important information. This would help in developing water quality and sediment quality guidelines for all rice pesticides. 5. A trial needs to be established in a typical rice growing season to test the suitability of all the techniques developed and validated during this project. A future trail should focus on an integrated approach where ELISA, GC/MS, biomarker approach and whole sediment bioassays developed for this project should be conducted concurrently. The use of multiple lines of evidence would help the irrigation companies and regulatory authorities to manage and minimise the off-site migration of rice pesticides. These approaches can be used as a compliance tool by regulatory authorities.

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7. References Coleambally Irrigation Co-Op Ltd. Annual Environmental Report (2003) http://env.colyirr.com.au/AER2003/ Accessed 2/7/08); http://env.colyirr.com.au/AER2003/Chapters/Molinate%20results%20ColeamballyDIPNR%20report.pdf Ellman, G., Courtney, K., Andres, J., Featherstone, R.1961. A new and colorimetric determination of acetylcholinesterase activity. Biochem Pharmacol. 7, 88–795. Hyne, R and Aistrop, M. (2006). IREC FARMERS NEWSLETTER No. 171, Summer 2006. Kralj, M. B., P. Trebse and M. Franko. 2006. Oxidation as a Pre-step in Determination of Organophosphorus Compounds by the AChE-TLS Bioassay. Acta Chim. Solv. 53: 43-51. Lowry, O., Rosebrough, N., Farr, A., Randall, R., 1951. Protein measurements with the folin phenol reagent. J Biol. Chem. 193, 265–275.

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8. Appendices Appendix 1

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Rapid Response Screening Tools to Minimise Offsite Impacts of Rice Pesticides RIRDC Publication No. 09/148 By Anu Kumar, Wendy C. Quayle, Mark Stevens, Marianne Woods and Rai Kookana

In recent years, increasing awareness of the potential adverse effects of pesticides on the environment has resulted in greater public pressure to assess, monitor and minimise off-site impacts. The application of pesticides for pest control in ricebased cropping systems is commonly practised in Australia.

The Rural Industries Research and Development Corporation (RIRDC) manages and funds priority research and translates results into practical outcomes for industry.

Our business is about developing a more profitable, dynamic and sustainable rural sector. Most of the information we This report presents the development of rapid response tools produce can be downloaded for free or purchased from our for the purposes of integrated pesticide management leading to website: www.rirdc.gov.au, or by phoning 1300 634 313 (local the environmental sustainability of rice-production systems. call charge applies).

Most RIRDC books can be freely downloaded or purchased from www.rirdc.gov.au or by phoning 1300 634 313 (local call charge applies).

www.rirdc.gov.au

Contact RIRDC: Level 2 15 National Circuit Barton ACT 2600 PO Box 4776 Kingston ACT 2604 Ph: 02 6271 4100 Fax: 02 6271 4199 Email: [email protected] web: www.rirdc.gov.au