Monitoring and Sampling Manual 2009 · Monitoring and Sampling Manual 2009 ... 3.3.2 Preservation and storage ... Appendix H7 National taxonomic codes for macroinvertebrate families

Monitoring and Sampling Manual 2009 (Version 2, published June 2010)

Monitoring and Sampling Manual 2009 Environmental Protection (Water) Policy 2009

Version 2 September 2010 (July 2013 format edits)

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Prepared by: Department of Environment and Heritage Protection © State of Queensland, 2013. The Queensland Government supports and encourages the dissemination and exchange of its information. The copyright in this publication is licensed under a Creative Commons Attribution 3.0 Australia (CC BY) licence.

Under this licence you are free, without having to seek our permission, to use this publication in accordance with the licence terms. You must keep intact the copyright notice and attribute the State of Queensland as the source of the publication. For more information on this licence, visit http://creativecommons.org/licenses/by/3.0/au/deed.en ISBN 978-0-9806986-1-9 Disclaimer This document has been prepared with all due diligence and care, based on the best available information at the time of publication. The department holds no responsibility for any errors or omissions within this document. Any decisions made by other parties based on this document are solely the responsibility of those parties. If you need to access this document in a language other than English, please call the Translating and Interpreting Service (TIS National) on 131 450 and ask them to telephone Library Services on +61 7 3170 5470. This publication can be made available in an alternative format (e.g. large print or audiotape) on request for people with vision impairment; phone +61 7 3170 5470 or email <[email protected]>. Citation Department of Environment and Heritage Protection (2009) Monitoring and Sampling Manual 2009, Version 2, July 2013 format edits. This document contains the common techniques, methods and standards for sample collection, handling and data management for use by Queensland Government agencies, relevant persons and other organisations for release and impact monitoring and to assess the condition and trend of Queensland waters. July 2013

http://creativecommons.org/licenses/by/3.0/au/deed.en�

Contents 1 Introduction............................................................................................................................................................2

1.1 Edition identification.........................................................................................................................................2 1.2 Purpose of the manual.....................................................................................................................................2 1.3 Status of the manual........................................................................................................................................3 1.4 Intended users .................................................................................................................................................3 1.5 Content of the manual .....................................................................................................................................3 1.6 Limitations........................................................................................................................................................4 1.7 Disclaimer ........................................................................................................................................................4

2 PART A Sampling design overview.......................................................................................................................5 2.1 Sound sampling design is essential ................................................................................................................5 2.2 Determining the scope of the sampling strategy .............................................................................................7

2.2.1 Defining the aims and objectives of sampling.........................................................................................7 2.2.2 Define the spatial boundaries of sampling..............................................................................................7 2.2.3 Define the temporal scale of sampling....................................................................................................8 2.2.4 Define the frequency of sampling ...........................................................................................................8

2.3 Sampling design ..............................................................................................................................................8 2.3.1 Importance of understanding the system being sampled .......................................................................8 2.3.2 What to sample .....................................................................................................................................11 2.3.3 Where to sample ...................................................................................................................................12 2.3.4 When to sample ....................................................................................................................................12 2.3.5 How to sample ......................................................................................................................................13 2.3.6 Quality control in sampling....................................................................................................................14 2.3.7 Cost effectiveness.................................................................................................................................14

2.4 Transport and security of samples ................................................................................................................14 2.5 Contact laboratories.......................................................................................................................................15 2.6 Sampling schedule ........................................................................................................................................15 2.7 Useful source documents for sampling design..............................................................................................15

3 PART B Sampling physico-chemical indicators of water quality and environmental health ...............................17 3.1 Sampling in the field ......................................................................................................................................17

3.1.1 Using intermediate containers and sampling rods................................................................................17 3.1.2 Automatic samplers...............................................................................................................................18 3.1.3 Field filtration equipment.......................................................................................................................20 3.1.4 Items for sample security ......................................................................................................................20 3.1.5 Sample carrier boxes ............................................................................................................................20 3.1.6 Marking pens.........................................................................................................................................21 3.1.7 Camera .................................................................................................................................................21 3.1.8 Voice recorder.......................................................................................................................................21 3.1.9 Global positioning system (GPS) ..........................................................................................................21

3.2 Labelling.........................................................................................................................................................21

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3.3 Sample containers and preservation methods ..............................................................................................22 3.3.1 Sample containers ................................................................................................................................22 3.3.2 Preservation and storage......................................................................................................................23

3.4 Preventing contamination ..............................................................................................................................24 3.5 Collecting samples.........................................................................................................................................24

3.5.1 Surface waters ......................................................................................................................................25 3.5.2 Groundwaters........................................................................................................................................27 3.5.3 Sediments .............................................................................................................................................27 3.5.4 Fish and other aquatic animals .............................................................................................................27 3.5.5 Vegetation and algae ............................................................................................................................28 3.5.6 Wipe sampling of surface contaminants (also known as ‘swab sampling’) ..........................................29

3.6 Instrument-based field tests...........................................................................................................................29 3.6.1 General guidance on taking field measurements .................................................................................30 3.6.2 Overview of field measurements...........................................................................................................32 3.6.3 Test kits.................................................................................................................................................35

3.7 Sample security and transport .......................................................................................................................36 3.7.1 Securing your samples..........................................................................................................................36 3.7.2 Transporting your samples ...................................................................................................................37

3.8 Laboratory analysis........................................................................................................................................40 3.8.1 Selection of analytical methods ............................................................................................................40

3.9 Data analysis and interpretation ....................................................................................................................41 3.9.1 Sources of reference values .................................................................................................................41

3.10 Data custodianship, management, and submission for regulatory purposes ...........................................42 Part C Appendixes......................................................................................................................................................43

Appendix C1 Forms ................................................................................................................................................44 Appendix C2 Methods for overcoming limit of detection problems: in situ extractions and the use of passive samplers..................................................................................................................................................................45

C2.1 In situ extraction ....................................................................................................................................45 C2.2 Passive sampling devices.....................................................................................................................46

Appendix C3 Flow measurement............................................................................................................................49 Appendix C4 Sampling water quality in temporary waters .....................................................................................49

C4.1 Sampling the receiving environment.....................................................................................................49 C4.2 Sampling release waters.......................................................................................................................50 C4.3 What to sample .....................................................................................................................................50

Appendix C5 Bulk natural water and sediment collection for direct toxicity assessment (DTA).............................52 Appendix C6 Contact details for laboratories..........................................................................................................53 Appendix C7 Units and concentrations...................................................................................................................54 Appendix C8 Sample containers and preservation methods..................................................................................58 Appendix C9 Fluvial sediment sampling using P 61 sediment samplers and Helley–Smith bedload samplers.....69

C9.1 Skills/competency and experience........................................................................................................69 C9.2 Equipment .............................................................................................................................................69 C9.3 Method ..................................................................................................................................................71

C9.4 Equipment use ......................................................................................................................................75 C9.5 Equipment maintenance .......................................................................................................................82 C9.6 Sample handling ...................................................................................................................................82 C9.7 Quality assurance .................................................................................................................................82 C9.8 References............................................................................................................................................83

Appendix C10 Sampling procedures for suspended solids and nutrients—application of water sampling technique.................................................................................................................................................................84

C10.1 Background for sampling procedures .......................................................................................................84 C10.2 Manual sampling procedures....................................................................................................................84 C10.3 Automatic sampling procedures ...............................................................................................................85 C10.2 Sampling procedures for filtered nutrients ................................................................................................86

PART D Sampling bio-indicators of water quality and environmental health .............................................................88 4.1 Macro-invertebrate sampling and assessment..............................................................................................88

4.1.1 Introduction to AusRivAS ......................................................................................................................88 4.1.2 Sampling program.................................................................................................................................88 4.1.3 Site selection.........................................................................................................................................89 4.1.4 Sampling frequency ..............................................................................................................................90 4.1.5 Habitats sampled ..................................................................................................................................91 4.1.6 Preparing for a field trip.........................................................................................................................93 4.1.7 Field sheets...........................................................................................................................................93 4.1.8 Water quality sampling..........................................................................................................................94 4.1.9 Biological sampling ...............................................................................................................................94 4.1.10 Laboratory macroinvertebrate sample processing ..................................................................................96 4.1.11 Database entry and software support ...................................................................................................98

4.2 Blue-green algae (cyanobacteria) sampling and assessment.......................................................................99 4.2.1 Introduction to blue-green algae ...........................................................................................................99 4.2.2 Guidelines for assessing blue-green algae...........................................................................................99 4.2.3 Monitoring sites ...................................................................................................................................101 4.2.4 Equipment required for algae sampling ..............................................................................................102 4.2.5 Sample collection ................................................................................................................................102 4.2.6 Contingency plan framework for blue-green algae response .............................................................105

4.3 Sampling fish ...............................................................................................................................................107 4.3.1 General considerations .......................................................................................................................107 4.3.2 Sampling fish using drift nets ..............................................................................................................108 4.3.3 Sampling fish using tow nets ..............................................................................................................108 4.3.4 Sampling fish using short seine nets ..................................................................................................109 4.3.5 Sampling fish using long seine nets....................................................................................................110 4.3.6 Sampling fish using fyke nets .............................................................................................................111 4.3.7 Sampling fish using cast nets .............................................................................................................114 4.3.8 Sampling fish using gill nets................................................................................................................116 4.3.9 Baited trap fishing.....................................................................................................................................117 4.3.10 Sampling fish using electrofishing ..........................................................................................................117

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4.4 References...................................................................................................................................................123 Part E Preparation of aquatic animal tissues (fish and crustaceans) for veterinary laboratory examination ...........124

5.1 Reasons for sending aquatic animal tissues for veterinary laboratory examination ...................................124 5.2 Collecting finfish specimens for diagnostic laboratory examination........................................................125

5.2.1 Sampling live finfish ............................................................................................................................125 5.2.2 Sampling and preparing fixed finfish specimens ................................................................................125 5.2.3 Freshly killed finfish on ice ..................................................................................................................125 5.2.4 Frozen finfish.......................................................................................................................................126 5.2.5 Fixatives and anaesthetics..................................................................................................................126 5.2.6 Basic anatomy of finfish ......................................................................................................................127 5.2.7 Gill and skin smears and wet mounts .................................................................................................127 5.2.8 Finfish dissection.................................................................................................................................129

5.3 Collecting crustacean specimens for diagnostic laboratory examination................................................130 5.3.1 Sampling live crustacea ......................................................................................................................130 5.3.2 Sampling fixed specimens ..................................................................................................................130 5.3.3 Basic anatomy of crustacea................................................................................................................131 5.3.4 Sampling and preparing fixed crustacean specimens ........................................................................132 5.3.5 Gill, appendage and larval wet mount preparations ...........................................................................134 5.3.6 Sampling and preparing molluscs.......................................................................................................135

Part F Monitoring mangrove forest health ................................................................................................................139 6.1 Mangrove litter trapping...........................................................................................................................139

6.1.1 Introduction—what is litter and how is it related to mangrove forest health? .....................................139 6.1.2 Why monitor litter productivity?...........................................................................................................139 6.1.3 Method summary ................................................................................................................................139 6.1.4 Site selection.......................................................................................................................................140 6.1.5 Installing litter traps .............................................................................................................................140 6.1.6 Emptying the traps ..............................................................................................................................140 6.1.7 Sorting trap contents ...........................................................................................................................140 6.1.8 Dry and weigh trap contents ...............................................................................................................141 6.1.9 Data interpretation...............................................................................................................................142 6.1.10 References and further reading..............................................................................................................142

6.2 Seedling regeneration .............................................................................................................................142 6.2.1 Introduction—why monitor mangrove seedlings?...............................................................................142 6.2.2 Method summary ................................................................................................................................143 6.2.3 Site selection.......................................................................................................................................143 6.2.4 Establishing a belt transect .................................................................................................................143 6.2.5 Tag seedlings......................................................................................................................................143 6.2.6 Measure height ...................................................................................................................................144 6.2.7 Measure stem diameter and stem density ..........................................................................................144 6.2.8 Count the leaves .................................................................................................................................144 6.2.9 Record sediment type, pH and salinity ...............................................................................................144

6.2.10 Draw a mud map of the site ...................................................................................................................144 6.2.11 Measurements needed during re-survey................................................................................................144 6.2.12 Data interpretation ..................................................................................................................................145 6.2.13 References and further reading..............................................................................................................145

6.3 Canopy cover and leaf area index...........................................................................................................145 6.3.1 Introduction—what is leaf area index and how is it related to mangrove forest health? ....................145 6.3.2 Method summary ................................................................................................................................146 6.3.3 Site selection.......................................................................................................................................146 6.3.4 Using the light meter ...........................................................................................................................146 6.3.5 Data interpretation...............................................................................................................................148 6.3.6 References and further reading ..........................................................................................................149

6.4 Mangrove forest structure .......................................................................................................................149 6.4.1 Introduction—what is mangrove forest structure and how is it related to mangrove forest health?...149 6.4.2 Method summary ................................................................................................................................149 6.4.3 Site selection.......................................................................................................................................149 6.4.4 Lay out transect and set up quadrats..................................................................................................149 6.4.5 Estimate canopy cover........................................................................................................................151 6.4.6 Estimate canopy dominance...............................................................................................................152 6.4.7 Measure stem diameter ......................................................................................................................152 6.4.8 Count saplings and seedlings .............................................................................................................153 6.4.9 Estimate height ...................................................................................................................................153 6.4.10 Soils ........................................................................................................................................................154 6.4.11 Tag and record position of trees.............................................................................................................154 6.4.12 Measurements needed during re-survey................................................................................................154 6.4.13 Data interpretation ..................................................................................................................................154 6.4.14 References and further reading..............................................................................................................156

6.5 Crab burrow counts .................................................................................................................................156 6.5.1 Introduction—why monitor crab hole density?....................................................................................156 6.5.2 Method summary ................................................................................................................................156 6.5.3 Site selection.......................................................................................................................................156 6.5.4 Establish transects ..............................................................................................................................157 6.5.5 Count the number of crab burrows .....................................................................................................157 6.5.6 Data interpretation...............................................................................................................................157 6.5.7 References and further reading ..........................................................................................................157

Part G Monitoring seagrass ......................................................................................................................................158 7.1 Intertidal percentage cover......................................................................................................................158

7.1.1 Introduction .........................................................................................................................................158 7.1.2 What is seagrass percentage cover?..................................................................................................158

7.2 Method for measuring seagrass percentage cover.................................................................................158 7.2.1 Method summary ................................................................................................................................158 7.2.2 Site selection.......................................................................................................................................159

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7.3 Data interpretation...................................................................................................................................161 7.4 References and further reading...............................................................................................................161

Appendixes ...............................................................................................................................................................162 Appendix H1 Checklist of equipment needed for macroinvertebrate field sampling ............................................162 Appendix H2 Queensland Site Information Sheet ................................................................................................163 Appendix H3 Water Quality Sampling Field Sheet ...............................................................................................168 Appendix H4 Field Sheets.....................................................................................................................................169 Appendix H5 Keys used for identification of Queensland Macroinvertebrate Fauna ...........................................170 Appendix H6 List of predictor variables used for the Mark I and Mark II models .................................................173 Appendix H7 National taxonomic codes for macroinvertebrate families collected in Queensland.......................174 Appendix H8 Transport of live aquatic animals.....................................................................................................176

Glossary....................................................................................................................................................................177 References................................................................................................................................................................185

Preface Water monitoring is undertaken by the Queensland Government for a variety of reasons, including the provision of information to government for policy and investment decision-making, to underpin natural resource management decisions by government and stakeholders, to assess impacts on the environment and to educate and inform stakeholders and the community generally.

Monitoring is also required to be conducted by persons under statutory approvals, and is additionally conducted by other organisations across Queensland, including local government, industry, regional natural resource management bodies and community groups. Many of these organisations collect valuable information on the condition of Queensland waters that complement Queensland Government monitoring.

The Monitoring and Sampling Manual 2009 provides the common techniques, methods and standards for sample collection, handling and data management for use by Queensland Government agencies, relevant persons and other organisations.

Where monitoring is required under legislation to be carried out under a protocol, the Monitoring and Sampling Manual 2009 is the principal document to decide the protocols. This manual is intended to be used by persons and organisations involved in the monitoring of the condition and trend of Queensland waters.

The Monitoring and Sampling Manual 2009 will facilitate consistency and increased scientific rigour of monitoring data available for interpretation by all stakeholders. It will allow them to assess the condition and trend of Queensland waters so that the aquatic environment can be managed for sustainable development and aquatic ecosystem health.

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1 Introduction

1.1 Edition identification This second edition of the Monitoring and Sampling Manual 2009 (the manual) supersedes the first edition of that manual, and sampling manuals published by the former Environmental Protection Agency, Department of Primary Industries, and Department of Natural Resources and Mines.

1.2 Purpose of the manual The purpose of the manual is to provide the common techniques, methods and standards for sample collection, handling, quality assurance and control, custodianship and data management, for use by Queensland Government agencies, relevant persons and other organisations.

The manual is a part of an integrated monitoring framework to decide the priorities, indicator selection, data storage, data analysis and reporting, as shown in Figure 1 below.

Where monitoring is required under legislation to be done under a protocol, including the Environmental Protection (Water) Policy 2009 (EPP (Water)) and the Environmental Protection Regulation 2008, the manual is the primary document to decide the protocols.

Figure 1.1 Integrated monitoring framework

1.3 Status of the manual This manual is the updated version of the primary document originally listed in the Environmental Protection (Water) Policy 1997 for use in deciding ‘protocols’. Accordingly, if there is any inconsistency between this manual and the other documents, this manual takes precedence to the extent of the inconsistency.

Some standards and other documents that have been used in preparing this manual will be revised from time to time. This could result in a procedure differing from that presented in the manual. The rules provided in Box 1.1 cater for such instances. In any situation where users of this manual use a revised document and/or adopt a protocol on the advice of an analyst, they should note the use of any revisions adopted and keep a record of the analyst’s advice.

Box 1.1 When a standard is revised

If this manual is found to differ from a revision of any of the other documents listed in the EPP (Water), amended after the date of this manual, a user may decide that the updated version of the document is more appropriate than the manual for deciding a ‘protocol’, either generally or in particular circumstances. This manual should not be interpreted as being inconsistent with the other document provided that:

• the revision of the document is subsequent to the date of this manual

• the specific procedure in the document is revised in the updated document

• an analyst advises that use of the revision in deciding the ‘protocol’ will lead to a determination of a quality as good as or better than that derived from using the manual.

1.4 Intended users This manual is intended to be used by:

• those who hold instruments under the Environmental Protection Act 1994 such as Environmental Authorities and Development Approvals (that comprise licences and approvals to carry out environmentally relevant activities) and Environmental Protection Orders

• employees and consultants of those who hold instruments under the Environmental Protection Act 1994

• those who analyse the samples collected for water quality determinations

• other persons and organisations involved in the monitoring of the condition and trend of Queensland waters.

1.5 Content of the manual The manual’s content has been prepared in consultation with other Australian environmental agencies and Queensland state and local government agencies. Australian and New Zealand and relevant international standards were considered during the preparation of this edition.

The manual is consistent with the nationally adopted framework presented in the Australian Guidelines for Water Quality Monitoring and Reporting (ANZECC/ARMCANZ 2000) and covers the sections of the framework indicated in Figure 1.2 below.

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Figure 1.2 Framework for a water quality monitoring program—Australian Guidelines for Water Quality Monitoring and Reporting (2000).

This manual presents procedures for:

• sampling design

• sampling in the field:

o making in situ tests and water quality measurements

o taking samples for water quality assessments, including samples of wastewaters, environmental waters, sediments and biota

o preserving and storing samples for water quality assessments, including samples of wastewaters, environmental waters, sediments and biota

o security and transport of samples

• arranging laboratory analysis

• data analysis and interpretation.

1.6 Limitations The manual cannot cover every set of circumstances encountered when determining a ‘protocol’ for sampling, and may not always provide sufficient or relevant directions. In situations where the user has little confidence that the samples might produce useful data, this should not stop them from being collected, particularly if there is no other opportunity to obtain the information they could provide.

1.7 Disclaimer Where particular brand names of equipment are mentioned, this has been done for illustration purposes only. Other makes or brands providing equivalent function are equally applicable.

2 PART A Sampling design overview

Sampling DesignSampling Design

Scope ofSamplingScope ofSampling

Why Sample?Why Sample?

SpatialBoundaries?

SpatialBoundaries?

Duration?Duration?

Frequency?Frequency?

Scope ofSamplingScope ofSampling


SpatialBoundaries?

SpatialBoundaries?

Duration?Duration?



SpatialBoundaries?

SpatialBoundaries?

Duration?Duration?


Sampling Design

Sampling Design

What to Sample?What to Sample?

Where to Sample?Where to Sample?

When to Sample?When to Sample?

How to Sample?How to Sample?

QCQC

CostEffectiveness

CostEffectiveness

What to Sample?What to Sample?

Where to Sample?Where to Sample?

When to Sample?When to Sample?

How to Sample?How to Sample?

QCQC

CostEffectiveness

CostEffectiveness

Transport & Security


Check Transport?Check Transport?

Securing SamplesSecuring Samples

DocumentationDocumentation









ContactLaboratories

ContactLaboratories

Double-checkRequirementsDouble-checkRequirements

Turn-around TimesTurn-around Times

Accreditation?Accreditation?

ContactLaboratories

ContactLaboratories







Figure 2.1 Essential components of a comprehensive sampling design

2.1 Sound sampling design is essential Before any sampling is carried out, the aim of the sampling exercise and how the results will be used need to be established. For example, a single result might be compared with a written specification or benchmark (such as release limits in an Environmental Authority or Development Approval or with relevant guidelines). Alternatively, several results might be used in calculations, and if so, it needs to be understood how the calculated quantities will be used. Prior to collecting samples, the need to assess the circumstances and the kinds of inferences that could be drawn from the results of sample analyses should be determined. That information will help identify where and when sampling should take place, and the quality characteristics that need to be determined for those samples.

The essential features of a sampling strategy are to ensure that the material sampled is genuinely representative of the body of material from which it was collected, that in situ measurements are reliable, and that the integrity of materials sent for laboratory analysis has not been compromised by contamination, degradation, transformation or losses.

The basic intent of environmental analysis is that analysis is carried out with selected portions (i.e. samples) from the location of interest, and the quality of the source material is then inferred from that of the samples. If the source material quality is temporally and spatially consistent then this inference would be uncomplicated. However, such constancy is rarely, if ever, observed in the real world. For example, virtually all waters show both temporal and spatial variations in quality (see Box 2.1 and Box 2.2), and consequently the timing and choice of location/s for taking water samples must be chosen with great care. Other materials such as sediments and biota typically also show such variations.

When designing monitoring programs it is important to ensure that the sampling regime is representative of the system and parameter/s of interest. For example, where a water body is well mixed and a parameter of interest is evenly distributed in the water column a grab sample may be appropriate. However, it is important to consider that parameters of interest may not be equally distributed. In such circumstances it may be necessary to assess the variability of the parameter of interest within the water column prior to sampling.

Such an evaluation would aim to determine whether sampling at a particular point is representative of a homogenous unit of water at a given point in time. Where the extent of variation is not known, the variation might need to be established by a pilot sampling program designed for that purpose, or a comprehensive range of

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samples taken to enable variability to be determined along with the primary aim of the sampling exercise.

In circumstances where undertaking an assessment of variability is not practical or possible it is recommended that information from relevant peer reviewed literature on the likely variability is used to provide guidance on an appropriate sampling strategy.

A similar approach to dealing with variability is recommended when designing sampling programs involving the collection of other materials such as sediments and biota.

When deciding the number of samples to collect and the frequency of sampling required, ideally, sufficient samples and replicates would be collected to represent the full range of variability present in space and time. Sampling designs should ultimately be defined by program objectives that can include the required statistical power required for discriminating between hypotheses or be based on the levels of acceptable sampling variability. Sampling designs should also be guided by the ’where to sample‘ and ’when to sample‘ sections in this manual (Sections 2.3.3 and 2.3.4).

Box 2.1 Variability of water quality over time

If the environment to be sampled shows changes over time—for example, river systems within minutes or hours, or lakes within days or weeks—the temporal pattern of sampling is of great importance. The schedule for the sampling program should take account of the expected temporal resolution of changes in the environment. In programs for monitoring wastewater treatment effluents, sampling around the clock may be required to determine whether control variables have been met or exceeded. A single sample can only be a snapshot at a single point in time and may not reliably represent typical conditions for a system that varies over time.

If many samples are taken over a period of time, it is often appropriate to match the sampling rate to the expected pattern of variation in the environment.

When it is necessary to quantify a contaminant load, multiple sampling periods may be needed. For example:

• Time-proportional sampling: samples containing identical volumes are taken at constant time intervals.

• Discharge-proportional sampling: the time intervals are constant but the volume of each sample is proportional to the volume of discharge during the specific time interval.

• Quantity-proportional sampling (or flow-weighted sampling): the volume of each sample is constant but the temporal resolution of sampling is proportional to the discharge.

• Event-controlled sampling: depends on a trigger signal (e.g. a discharge threshold). For example, to detect peak concentrations during short-term changes of water quality, event-controlled samplers are useful.

Alternatively, passive samplers can be used to integrate variations in water quality over an extended period of time. For further information on the applicability and use of passive samplers see Appendix C2.

Box 2.2 Variability of water quality in space

It is important to understand how natural processes in environmental waters can affect water quality characteristics, and to be aware that water bodies are not homogeneous within a cross sectional area or depth profile.

Water bodies can be stratified (layered). This means the composition of the different layers is substantially different in respect of at least one characteristic. For example, in estuaries, water quality characteristics can vary because of ingress/egress of saline waters. Estuaries are commonly stratified when freshwater flow is much larger than tidal flow; the fresh flows seawards over the saline waters and a ‘salt wedge’ develops. Stratification could also result from temperature effects in waters with low current velocities. Such stratification is usually most pronounced in summer months when surface waters are much warmer than bottom waters. After separation, the water layers often develop markedly different chemistry. Such layers also tend to prevent mixing of discharged contaminants.

When sampling environmental waters (typically, when investigating a pollution incident), it could be important to remember that stratification might have occurred, and to take measurements at different depths to show whether this is so. The reverse process (de-stratification) can occur when the seasons change. The resulting inversion (‘turnover’) of the water can result in low oxygen water rising to the surface and causing adverse effects (such as odours from anaerobic decomposition at depth, and/or nutrient/metal enrichment). Other examples include the distribution of suspended solids within the water column from physical processes of re-suspension, deposition and flocculation. The concentration of suspended solids is dynamic in the water column and can fluctuate naturally according to flow conditions and water chemistry.

When the purpose of sampling is to assess compliance with a statutory provision such as a condition attached to an Environmental Authority or Development Approval, the sample should be taken to provide a reliable measure of the specific characteristic or parameter specified (e.g. the concentration of suspended solids at a defined sampling point). Where the statutory provision is not explicit, the sample should represent fairly the body of material from which it is taken during the period of the sampling.

Where the aim of the sampling is to measure compliance with conditions of an environmental authority or development approval, and the conditions include a statistical sampling regime, this should be followed so that the results can be of use. However, if there is reason to believe variability is a confounding factor, additional samples may be needed to check this out.

The Environmental Protection Act 1994 and its subordinate legislation, including the Environmental Protection Regulation 1998 and the Environmental Protection (Water) Policy 2009, must be taken into account when deciding where and when to sample for compliance in a pollution investigation, checking compliance with an environmental authority or development approval, or undertaking a receiving environment monitoring program.

Reference should be made to the current conditions of any relevant licence or permits, particularly when confirming compliance. The conditions may include specific sampling locations, times of release and quality characteristics that will assist with designing the sampling strategy.

2.2 Determining the scope of the sampling strategy

2.2.1 Defining the aims and objectives of sampling

2.2.1.1 Purpose of sampling

Monitoring and assessment that involves sampling can be undertaken for a range of reasons. Some of the primary reasons covered by this manual include:

• investigating pollution incidents—sampling at a site where pollution has been reported can be challenging as you might not be aware of the constituent pollutants, or often the source of the release. In such cases, the aim of sampling should be to obtain evidence that will:

o discover and prove the nature, the source, and the effects of the contaminants

o be performed in such a way as to be legally admissible in court

• confirming compliance to licence conditions of an environmental authority or development approval—sampling of wastewater that is stored or released by the holder of an environmental authority or development approval or similar legal instrument is often performed routinely to confirm compliance with the imposed conditions of release. To test for compliance with the conditions, samples must be collected in a manner that will ensure valid analysis results for those particular contaminants

• undertaking a receiving environment monitoring program

• undertaking an environmental evaluation of an activity.

2.2.1.2 Specific sampling objectives

The objectives of the sampling program should be determined and documented. These should be as specific as possible. Common sampling objectives include:

• determining if one or more contaminants found in a waste or in the environment have originated from a singular or multiple source

• determining whether one or more contaminants in a release are in sufficient quantity to cause adverse environmental effects consistent with those observed at the time of the incident

• determining whether the contaminants in a waste release are having a measurable impact on the receiving environment water quality and whether environmental values are being affected

• determining whether the quality of waters have changed significantly, consistent with the definition of the term ‘environmental harm’ in the Environmental Protection Act 1994, and confirm whether the observed environmental change(s) occurred as a result of the release.

2.2.2 Define the spatial boundaries of sampling The geographic boundaries of the sampling event should be based on the issue of concern and the ecosystem type rather than on convenience and/or budgets. For example, some important considerations would include:

• the likely spatial uniformity of the parameter(s) of interest

• the size of the area to be assessed.

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2.2.3 Define the temporal scale of sampling Temporal scale refers to the length of time over which a system is to be observed; that is, the appropriate period of time over which the samples are to be collected.

Different processes operate at different temporal scales, and the sampling designer should incorporate all the important time-related considerations into the design. For example, the movement of sediment in a river system occurs over tens of years at the catchment scale, whereas toxicant effects may occur over days (transient) or be continuous in nature. The temporal scale and, similarly, the frequency of sampling (see below) need to be suited to the temporal characteristics and occurrence of the contaminant.

2.2.4 Define the frequency of sampling Consideration needs to be taken of the frequency of observations (sampling events) required to provide sufficient resolution of the issues of concern. Sampling may be required every hour, day, week, fortnight, month or possibly only once a year. The sampling designer needs to determine a frequency of sampling (level of resolution) that is sufficient to satisfy the requirements of the program objective, yet not sample too frequently and cost more than necessary.

2.3 Sampling design

2.3.1 Importance of understanding the system being sampled The achievement of good sampling design can be assisted if the designer has some understanding of the ecosystem for which the sampling program is being designed. This understanding is best formalised in a conceptual model (or process model) of the system being examined. The model can be a simple box diagram that illustrates the components and linkages in the system, or a graphical representation of the system. Whatever model is used, it should present the factors that are perceived to be driving the changes in the system and the consequences of changes to these factors.

During the formulation of a model, several decisions must be made or the model will be too complex. For example:

• What are the major issues of concern (e.g. nutrients, metal loads, bioavailable metals)?

• What ecosystem (including subsystem type) should the model describe

• (e.g. freshwater, marine waters, estuarine waters, wetland, seagrass bed, mangroves)?

• Which state of flow should the model describe (e.g. base flow, flood event)?

Once formulated, the process model can be used to help define:

• important components of the system and the important linkages

• key processes

• cause–effect relationships

• important questions to be addressed

• spatial boundaries

• valid measurement parameters for the processes of concern; what to measure, and with what precision

• site selection

• time and seasonal considerations.

Examples of graphical conceptual models that may assist sampling design are at Figures 2.2, 2.3 and 2.4.

The importance of having an understanding of the ecosystem for which the sampling program is being designed is demonstrated by the complexity of nutrient cycling processes in waterways.

Plants use light as a source of energy for everyday growth and repair. They also require elements such as carbon (which they derive from carbon dioxide in the atmosphere) and the nutrients nitrogen (N) and phosphorus (P). Nutrients stimulate the growth of aquatic plants and are required to maintain the productivity of ecosystems. However, the growth of aquatic plants can be limited, despite there being sufficient light available, when nutrients are present only at minimal concentrations. Nutrients can become an environmental problem when they occur at excessive concentrations. Negative effects of higher-than-normal nutrient concentrations include the eutrophication of waterways.

Figure 2.2 Conceptual diagram of a coastal system including anthropogenic activities, inputs to waterways and areas of value

Typical direct effects of eutrophication include increased frequency of algal blooms (including toxic algae) and hypoxia. Increased production of aquatic plants and algae may temporarily increase oxygen production within a water body, but when these decompose, this may cause a depletion of dissolved oxygen (DO). Low DO levels can lead to fish kills and the death of other aquatic fauna. Other effects of eutrophication may include increased turbidity and changes in community composition.

The nutrients N and P occur naturally in Australian surface water systems. They often occur in both particulate (i.e. organic and sediment-bound) and dissolved forms. The movement of nutrients from land may originate from both diffuse and point sources. Pathways for diffuse sources include riparian litter fall, soil erosion and sediment transport (see Figures 2.3 and 2.4). Fertilisers may be a source of N and P in agriculturally dominated catchments. The concentration and types of particulate and dissolved forms of N and P in waterways can indicate potential stresses from land uses and land management practices in the catchment.

An understanding of the relationship between flow and nutrient concentrations, coupled with N and P cycling (and the transformation from one form to another), is crucial when interpreting concentrations and loads and subsequently making any type of assumption or conclusion (e.g. is the nutrient a new contribution to the cycle or a transformation of a previously deposited load?). Without this understanding, changes in N and P concentrations and/or loads may be more attributable to variable flow regimes and biological factors rather than any imposed management action.

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Figure 2.3 Typical processes affecting nutrients within different parts of a catchment

Figure 2.4 Illustration depicting typical nutrient and sediment inputs into catchments

2.3.2 What to sample What material is relevant to collect samples of, and what measurements should be taken?

When checking compliance, environmental authority or development approval conditions typically specify the contaminants and the permitted ranges of concentrations allowed in the release. However, in cases of suspected environmental pollution incidents, you might not know what pollutants are present. Furthermore, when sampling the receiving environment, there are a range of related indicators that may need to be measured across different media such as surface water, sediments or biota. This section provides guidance on determining which characteristics should be sampled.

When choosing indicators, it is important to know whether there are defined benchmarks such as water quality objectives, guidelines, limits or other standards relevant to your situation to compare with your measured data. Indicators may be chosen because they have such benchmarks and may best indicate water condition or potential environmental harm. If no defined benchmarks exist, it is essential that your sampling design includes appropriate reference or control sites so that you are able to make a comparison against something.

2.3.2.1 Sampling media

The aim of sampling is to estimate quality characteristics of one or more of the following:

• wastes released to a water body (or potentially released)

• the receiving environment via:

o a permanent or temporary water body—usually surface waters, but occasionally groundwater

o bottom sediments of a permanent or temporary water body

• specimens of animal or plant life thought to have been affected by a release or by a change in natural conditions.

2.3.2.2 Sampling waste streams

Environmental authority or development approval conditions typically specify the contaminants and the permitted ranges of concentrations allowed in the release. Additional characteristics may provide greater information about the potential environmental harm that might be caused, for example, although only biochemical oxygen demand (BOD) might be specified in the environmental authority or development approval, chemical oxygen demand (COD) and total organic carbon (TOC) often provide more information, and could be worth assessing. It may also be important to measure other characteristics due to a change in an operating condition or a specific incident.

The characteristics being measured should relate to the potential contaminants used and/or generated in the process that produces the waste stream, in addition to their potential effects on the receiving environment.

In addition to measuring water quality characteristics, flow measurement of wastes/wastewater is often required for point source releases. This allows regulation and quantification of flow and loads of contaminants. Flow measurements of water bodies can be important for regulation as they can be used to assess initial mixing of point source discharges or as triggers to allow licensed discharges, particularly for event-based releases. Flow measurements of waterways may also be required for pollution incidents to assess or predict the extent of impact. Further information on flow measurement can be found in Appendix C3.

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2.3.2.3 Sampling the receiving environment

Your assessment should take into account:

• potential sources of contamination

• likely contaminants

• type of waterway and flow rates; whether freshwater, estuarine or marine, and whether a flowing stream, lake, or ephemeral (in which case it may be wet or dry, or evaporating and concentrating contaminants at the time of sampling)

• licensed releases into the waters

• potential sources of releases

• recent weather such as heavy rain, showers or drought conditions

• historical occurrences of similar incidents.

2.3.3 Where to sample

2.3.3.1 Where should samples be collected and measurements taken?

Many environmental authorities have conditions which specify where samples are to be taken. Some have more than one sample point (two or more outlets, or an intake as well as an outlet). Where no sampling location is specified, you should take a sample from a site you judge to be representative of the release material (and the receiving waters, where relevant).

In more populated areas identification of a particular location can be assisted by a map of the area at a suitable scale (typically 1:25 000 or larger), or an aerial photograph showing individual buildings, preferably printed beforehand and taken with you. In more remote areas, use other points of reference, such as topographic features (hills, quarries, stream bends, etc.) or structures like fences or stone walls, and a global positioning system (GPS).

When investigating environmental pollution incidents, you should consider all possible sources of the pollutant, including licensed and unlicensed sources of release. You should aim to sample at the site of the pollution reported, at the point of any suspected contributing releases and also in an area upstream/distant from the suspected source.

It is important to identify with sufficient accuracy the location from which a sample has been collected to avoid raising a doubt about ‘what the sample represents’, particularly in cases where a location a few metres away might have given significantly different results. For example, in a river, was the location upstream or downstream of a tributary stream; or on the inside or outside of the bend? Or, where multiple discharge points exist, could it have been the one ‘next to’ the one claimed? This last instance could be significant when investigating complaints in situations where you do not know what other discharges may have occurred.

2.3.4 When to sample

2.3.4.1 Timing of sampling

The timing of the sampling should address the pre-defined objectives of the program. When there is a suspected environmental incident, it is prudent to sample as soon as possible after the incident has occurred.

Some conditions on environmental authorities specify that release is to take place only at certain times of the day (for example, on an outgoing tide) or under certain weather conditions. This should be considered in your sampling design where applicable.

For impact assessments, sampling before and after is important (but not always possible), preferably with multiple before and after reference sites. In situations where there is no ‘before’ information available at the impact location, data collected by sampling from reference sites may be indicative of conditions at the impact location prior to the incident. For example, a chemical spill may have contaminated the receiving environment, and caused impacts on local biota, but there are no pre-spill data available. However, concentrations of contaminants or macroinvertebrate population indices measured at unimpacted reference sites after a chemical spill can be indicative of what those parameters could have been at the incident location prior to the spill.

Water quality varies with stream flow conditions, so in considering the timing of sampling, it is important to establish whether sampling during baseflow or during flood event conditions (or both) is appropriate.

2.3.4.2 Sampling during baseflow and event conditions

As maximum and minimum values for water quality indicators may be reached either during high and or low flow events, it may be necessary to target either baseflow and or event conditions of rivers or streams. Fluctuations in water quality may also occur due to effluent discharge from a point source. Baseflow sampling is carried out when the flow is predominantly influenced by groundwater and has little overland flow component (i.e. no flow derived from runoff after rainfall). Baseflow sampling of streams should be undertaken with adequate lag time following an event. In general, a period of at least six weeks following a major flow event should be allowed to elapse to ensure samples are solely baseflow water and are not capturing the tail of an event. An ’event‘ is classified as any significant rise in water level caused by rainfall. Sampling regimes for event sampling may be targeted at obtaining results from either smaller more frequent events, or larger but less frequent events. Episodic events are known to transport large quantities of some contaminants though, in some circumstances significantly higher concentration peaks can occur in much smaller but frequent flood events (see example in Figure 2.5). In some cases, these smaller more frequent events may also have a greater contribution to total or annual contaminant loads. Sampling of a flood event can include a series of discrete ’grab‘ samples collected to represent the extent of the event. See Appendix C10 for more information on grab sampling procedures.

Figure 2.5 Example of contaminant fluctuations with stream flow

2.3.5 How to sample

2.3.5.1 How many samples should be collected?

Unless the material being sampled is known to be well mixed (well mixed water body or end-of-pipe discharge), it is unlikely for a single measure to be representative of the source body of material. Multiple measurements are needed to allow the calculation of a mean and confidence interval for the characteristic of interest, or to allow statistical testing for significant differences between locations or non-compliance with statutory provisions. This requires multiple readings for in situ measurements and multiple samples where laboratory analysis is involved. A minimum is three data points per site for basic statistical tests, but more may be required depending on the inherent variability in the measurement data. Just how many data points are needed may not be known until after chemical analysis of some samples. Accordingly, it is sometimes good practice to take additional samples and to store these for subsequent analysis if required. However, the requirements of maximum holding times for many contaminants may make this untenable.

2.3.5.2 Is grab sampling adequate or should composite samples be taken?

Most samples taken will be grab samples—taken by filling sample containers over a ‘short’ period (seconds or minutes). A single grab sample may be used where a hazardous situation has arisen or is suspected and the sample is taken to confirm the presence of the hazardous substance.

A single grab sample may also be used where the body of water being tested is well mixed and its quality can be adequately described by a single sample. However, in many situations, a single grab sample in isolation is of limited use because it takes no account of variations in quality with time or space (see Box 2.1 and Box 2.2). In

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such a situation, the taking of a composite sample is a useful strategy. A composite sample may be temporal by combining contributions of material collected over a longer period (minutes, hours or days). Alternatively, a composite sample may be spatial, for example, comprising a series of equal contributions of material taken along a transect (e.g. across a channel). This gives a spatially ‘more representative’ sample than a single grab sample at a single point. An example of the advantage of using composite sampling occurs in measuring concentrations of total nitrogen from a sewage treatment plant for the purpose of estimating loads. A single weekly grab sample will not capture the variations across a day or week. A suitable alternative would be to take a 24-hour composite sample.

Notes:

Composite sampling will not provide information on the maximum concentration reached (i.e. spikes in concentrations) which is often relevant when dealing with toxicants.

The use of automatic samplers to prepare composite samples over a period of time could be problematic due to delays in delivering samples to the laboratory for analysis. Under some circumstances this might not be significant, but you should check with the analyst before sampling with such equipment.

2.3.6 Quality control in sampling Quality control is an important part of any sampling exercise. The purpose of a quality control scheme is to check whether bias, sample contamination, or analyte loss could affect the results, and so invalidate the process.

Suitable techniques include:

• reference sites: comparable (but unimpacted) locations where samples are taken for comparison with others (for example, upstream of a discharge point, or a tributary other than the one of interest). Unless the condition of the reference site/s is known, and because of variability, it is often wise not to rely on a single reference site

• control samples, including:

o field spikes—an uncontaminated sample of the media (e.g. water) contained and preserved in an identical fashion to the field samples is spiked with contaminants of interest and accompanies the field samples during the sampling. Analysis of the spiked sample quantifies analyte loss (if any) through comparison with the spiked concentration

o field, transport and container blanks—uncontaminated samples of the media (e.g. water) contained, handled (for example filtered) and preserved in an identical fashion to the field samples are analysed and measures of introduced contaminant (if any) are used to quantify and trace contamination problems associated with the sampling methods and materials.

These techniques are discussed in AS/NZS 5667.1:1998.

It is recommended that you discuss this aspect with the analyst during planning of a sampling exercise, particularly if the results are liable to be used in making decisions which could have significant health related, financial or legal implications.

2.3.7 Cost effectiveness It is preferable for the cost of sampling programs to be as small as possible while still meeting the stated objectives of the monitoring study. Cost-effectiveness considerations involve trade-offs between loss of statistical ‘power’ (i.e. the capacity of a program to discriminate between various hypotheses) and the cost of data acquisition. Costs of data acquisition taken into account for cost effectiveness include:

• the number of sampling stations, sampling occasions and replicates

• the cost of collecting samples (staff, transport, consumables)

• the cost of analysis

• the cost of data handling and interpretation (cost of reporting).

Cost-savings can result from collaborative monitoring, for example, when local councils pool resources with other water managers to comprehensively monitor a particular water body.

2.4 Transport and security of samples Samples collected in areas remote from the laboratory might need to be freighted by private companies—road or air transport—if the sampler cannot deliver them personally. The samples need to be delivered within the maximum holding times.

2.5 Contact laboratories When possible, contact the appropriate laboratories before going to the field to ensure that analysis can be performed before expiry of the maximum holding times. You should inform the laboratory of details concerning the samples you will be sending. One way is to send a completed copy of a form such as the Notice of Samples Expected shown in Appendix C1. Send this by mail when the sampling is planned some days ahead, or by facsimile or email if urgent. If this is impracticable, try to contact the analyst by telephone to give details of the samples.

You should also try to give the laboratory any information you can about the sample source, likely range of concentrations and purpose for which the results are to be used. This will help the analyst choose a suitable analytical method with appropriate ‘limit of reporting’ (LOR). In some cases the LOR can be improved if the analyst knows these details beforehand.

If sampling from areas affected by a chemical spill, try to advise the laboratory to expect high concentrations in the samples. Providing the laboratory with such information may help avoid results such as ‘out-of-range’ concentrations, and subsequent delays while your samples are diluted and re-analysed.

2.6 Sampling schedule Once the sampling design has been finalised a sampling schedule should be prepared that includes such information as:

• where and when the samples are to be collected

• source of each sample—whether from wastes, waters or sediments

• the nature of the material to be sampled

• the quality characteristics being sampled

• the sampling containers (and associated paraphernalia) needed

• preservatives needed

• the maximum holding time for each sample.

A blank copy of a sampling schedule form is located at Appendix C1. (Details of sample containers, preservatives and holding times are given in Appendix C8).

2.7 Useful source documents for sampling design Source documents for sampling design are shown in Table 2.1.

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Table 2.1 A selection of relevant Australian and New Zealand standards related to water and sediment sampling

AS/NZS 5667.1:1998 Water quality—Sampling

Part 1: Guidance on the design of sampling programs, sampling techniques and the preservation and handling of samples


Part 4: Guidance on sampling from lakes, natural and man-made


Part 5: Guidance on sampling of drinking water and water used for food and beverage processing


Part 6: Guidance on sampling of rivers and streams


Part 9: Guidance on sampling from marine waters


Part 10: Guidance on sampling of wastewaters


Part 11: Guidance on sampling of groundwaters


Part 12: Guidance on sampling of bottom sediments

AS/NZS 2031:2001 Selection of containers and preservation of water samples for microbiological analysis

AS2360:1993 various parts

AS/NZS 3778: various parts

Measurement of fluid flow in closed conduits

Measurement of water flow in open channels

3 PART B Sampling physico-chemical indicators of water quality and environmental health

3.1 Sampling in the field This section details the equipment and materials to be used in sampling. Appendix C1 contains a checklist of equipment and materials. You can copy this checklist for each sampling trip, and modify it as needed.

Sampling in the FieldSampling in the Field

Sample collection methods & equipment

Water, Sediments and/or Biota

Sample collection methods & equipment

Water, Sediments and/or Biota

Sample containersSample containers

Sample preservation & storageSample preservation & storage

Field measurementsField measurements

ConsiderQA/QC

ConsiderQA/QC

ConsiderOH&S

ConsiderOH&S

Figure 3.1 Necessary materials and equipment considerations for sampling in the field

3.1.1 Using intermediate containers and sampling rods It is always best to collect samples directly into the appropriate bottle or jar. However, it is sometimes necessary to use an intermediate container in order to collect samples, using buckets, beakers, pumps, filters, syringes, sediment grabs, trowels and/or sampling rods.

In using intermediate containers, contamination of the intermediate containers from previous samples becomes a risk. Contamination of the sample can occur by leaching of previous substances from the walls of the intermediate container, or loss of analyte by adsorption on to the walls of that container.

For samples containing substances in ‘trace’ concentrations, typically in the microgram-per-litre (µg/L) range (such as trace metals and pesticides), use of an intermediate container should be avoided if possible. Sometimes the use of ’intermediate containers‘ such as syringes and filtering equipment or components of automatic samplers is unavoidable, and so precautions to reduce the risk of contamination are critical.

If samples need to be collected using intermediate containers, it is important that intermediate containers be pre-washed (for example syringes and filters) or flushed (for example in the case of automatic sampler lines) with existing site water before being used for the final collection of samples. The use of field blanks (see section 2.3.6) can identify sample contamination issues.

A sampling pole with a large clamp (or other suitable device) to hold the sampling containers can be used to give greater reach when collecting samples (see Figure 3.2). If the rod becomes contaminated, wash it promptly, making sure the washings cannot contaminate any samples or any material about to be sampled (for example by disposing of washings downstream of the sampling site).

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Figure 3.2 Sampling pole and sample bottle

3.1.2 Automatic samplers In some circumstances it may be preferable to use an automated sampling device to collect samples (Figure 3.3). When sampling flood waters it is often unsafe to approach a stream bank to collect a sample manually due to the presence of high flows or in stormwater drains that can have flashy unpredictable flows. Also, in some instances it may be necessary to sample at regular intervals throughout a 24 hour time period or at times when it is not possible to collect them manually. In these situations it is appropriate that an automated sampler be used to collect grab samples. Automated sampling devices include refrigerated or non-refrigerated automatic pump samplers and rising/falling stage samplers.

Automatic pump samplers, both refrigerated and non-refrigerated are comprised of a number of bottles in a carousel, a sample intake line that is fixed in place within the stream and connected to a pump or pumps, and a computer controlled data logger that requires programming. The equipment is programmed to be ‘triggered’ when a pre-entered set of conditions are met. For example: a certain stream height, time of day, change in temperature, the rate of rise or fall of the stream level, a particular turbidity reading or any number of possible programmable triggers. Once triggered, the automatic sampler starts sampling according to the program set. Installation of automatic samplers should be in accordance with manufacturer instructions. When using automatic samplers for taking water samples, it is important to adhere to the requirements for sample handling including sample holding times and sample containers for the parameter of interest (see Appendix C8). If using automatic samplers, the sampling lines (tubing or pipes) must be regarded as an intermediate container (see section 3.1.1), and the possibility of exchange between the sample and the walls of the lines must be considered. The time of contact between the sample and the wall material is important, and residues of previous samples must be properly flushed out before liquid is delivered into the sample container. More detail on this aspect is given in AS/NZS 5667.1: 1998.

Automatic samplers also include remote samplers, such as rising-stage and falling-stage samplers. In regards to sampling event-based flows in ephemeral or temporary waterways, only falling-stage samplers should be considered, as rising-stage samplers tend to sample first flush waters. If the data is to be used to determine locally-derived water quality objectives (as per ANZECC/ARMCANZ 2000) then it is recommended that in accord with the AusRivAS sampling protocols (see section 4.1) sampling takes place 4–6 weeks after a flood event and flow has been established.

Figure 3.3 Auto-sampler

With rising stage samplers, water samples are taken as the river level rises and samples can only be retrieved after the river level has receded (Figure 3.4). As the water rises it reaches the crown of the intake, flow starts over the crown and begins to fill the bottle. Sample bottles fill progressively from bottom to top. Sampling ceases when the level of the water in the bottle reaches the inner end of the air exhaust, which then prevents circulation through the sampler. An air lock forms in the intake and prevents enrichment of the sample from water flowing back and forward and transporting sediment into the sample container.

Figure 3.4 Rising stage sampler

Rising stage samplers do not involve refrigeration and because samples are exposed to light and ambient temperatures they are only valid for total nutrient and sediment analysis. Note however that samples must be retrieved within a few hours for total nutrient analysis to be valid (particularly for N). Rising stage samplers are useful for collection of samples from flashy, intermittent streams at remote or sites that are not easily accessed.

Sampling units must be securely mounted, one above the other, with adequate support provided to prevent dislodging by large logs and other debris. Samplers should be erected and installed so that they are pointing in a downstream direction (ensure bottles are facing upstream). Sampler location is recommended in the following areas:

• on the inside of river bends (debris tend to be swept to the outside of bends)

• adjacent and mid stream of large trees (providing partial protection)

• downstream of small shrubs and trees (provide partial protection).

Each visit to the site should also involve inspecting sampling units for evidence of insects in the intake and breathing tubes, a common reason for missed samples (and contamination).

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3.1.3 Field filtration equipment For some characteristics listed in Appendix C8, the samples must be filtered in the field before they are placed in the container used for transport to the laboratory. Depending on the characteristics being measured, either vacuum or pressure filtration equipment may be needed.

An important factor in field filtration is to note that the equipment used for the filtering task is an ’intermediate container’ and as such the equipment is a potential source of contamination of the sample (see section 3.1.1) so that pre-rinsing and the use of field blanks (see section 2.3.6) is advisable.

Vacuum filtration is recommended for chlorophyll samples; the pressure difference across the filter should not exceed 40 cm of mercury to prevent lysis (rupturing) of the cells. You need to filter a known volume of the water and send only the filter paper to the analyst (see Appendix C8).The equipment comprises a vacuum flask, vacuum pump, sintered glass filter funnel and a supply of the specified filters. It is not essential to filter the volume shown in Appendix C8; usually, only one filter paper needs to be submitted. It is essential to record the actual amount of water passed through the filter and report this to the analyst, so that the concentration can be calculated.

Pressure filtration is recommended for dissolved metals and some nutrient characteristics. The water passing through the filter is sent to the laboratory and the filter discarded. Typical equipment comprises a hand-operated syringe with a supply of 0.45 µm filters and, for waters with high suspended particle load, pre-filters (see Figure 3.5).

Figure 3.5 Pressure filtration equipment

Occasionally, a high content of suspended matter could make it difficult to filter the volume shown in the table. In cases where the aim is to give the analyst a stated volume of filtered liquid, the use of more than one filter might solve the problem. Alternatively, you could measure or estimate the volume actually filtered, and provide this information when submitting the sample.

3.1.4 Items for sample security Ensure that you have an adequate supply of items such as sample seals, evidence bags, and locked storage boxes as detailed in section 3.7.1.

3.1.5 Sample carrier boxes To keep samples at suitably low temperatures they are transported in cleaned/ uncontaminated insulated carrier boxes (coolers). These are kept cool by adding block or crushed ice, dry ice, freezer-blocks, or other similar substance, or are refrigerated by a power source.

Samples requiring refrigeration are generally packed in crushed ice. Crushed ice is used in preference to block ice as it can be packed in much closer contact with the samples.

Note: An acceptable alternative to crushed ice is a combination of crushed ice and ‘ice bricks’ (frozen containers of refrigerant solution having a freezing point not greater than 0oC), provided that the ‘ice bricks’ are in a frozen state when the samples are packed in the carrier, are completely surrounded by crushed ice, and are used only to lessen the amount of crushed ice required—not as the sole or primary means of cooling the samples.

Dry ice (solid carbon dioxide) is used where samples must be frozen immediately after collection. It is available in block and pellet form. Pelletised dry ice is preferable for the same reason as for crushed ice. A combination of block and pellets could also be used, the pellets being placed next to the sample containers. Suppliers of dry ice are listed in the telephone directory or can be found via web search.

An acceptable alternative is a transportable freezer, provided that the samples, once placed in it and frozen, are

taken to the laboratory promptly, and care is taken to ensure that they do not thaw before delivery. In an emergency, samples can be frozen by surrounding them with a slurry of crushed ice mixed with common domestic salt (sodium chloride). This rapidly achieves temperatures well below 0oC.

When storing chilled or frozen samples in coolers, note that the coolers can be a source of sample contamination under some circumstances. For example, if a cooler has been used for storing fish, and is then used for storing samples collected for nitrogen or phosphorus, residual odorous substances from the fish (such as ammonia) can permeate the container walls, even if the container is of high density polyethylene (HDPE). If a cooler used for odorous material needs to be used for samples, be sure to clean it thoroughly before use.

If air transport of samples is involved, take account of air transport and company requirements regarding wet and dry ice.

Workplace health and safety—it is hazardous to transport dry ice inside a motor vehicle with all of the windows closed.

3.1.6 Marking pens Only waterproof pens should be used for labelling. Marking pens used to label samples must have waterproof ink. Enamel paint pens are useful in this regard. Writing a label before taking the sample (e.g. dipping the bottle into a waterway) avoids the difficulty of trying to write on a wet label. Only write on dry labels, and carry spare pens.

Note that when sampling waters for the presence of solvent-type compounds, extra caution should be used as marking pens contain solvents and could contaminate your sample.

It is advisable to place labelled samples inside a plastic bag to protect the label details from rubbing or scratching off.

3.1.7 Camera Photographs are useful when investigating pollution incidents. Check that you have appropriate spare items such as films, memory cards and batteries. Accessories such as glare/ultra-violet light filters, extra lenses or flash equipment could be appropriate in some circumstances.

Cameras that automatically imprint the exposure date and time on the photograph can be useful, but you need to be sure that the camera’s clock is set correctly. This also applies to digital cameras.

When taking photographs, it is essential to clearly identify when and where each shot is taken. Shots may be numbered consecutively and descriptive notes made at the time. Where practical, include identifying features such as a sample label or placard within each shot.

Note that conventional photographic images on film or paper, and electronic image files produced by scanning or by digital cameras, can all be manipulated. Therefore, images intended for legal purposes should be stored in secure places to guard against unauthorised access.

3.1.8 Voice recorder A voice recorder can be useful for immediate recording of your observations if circumstances make writing on paper difficult. If you use one, you should listen to the recording and transcribe it as soon as practicable.

3.1.9 Global positioning system (GPS) A hand-held GPS can be useful in quickly recording the location of your sampling sites by storing them as waypoints. Hand-held GPSs are typically accurate to around 5 m, but this can change with cloud cover, altitude, tree cover, the number of satellites the unit can ‘see’ at the time of the reading and which map datum you have used (e.g. Aust Geod. 84, WGS84, etc.). The accuracy of your readings, and which map datum you have used should be recorded at the time of your GPS reading.

If you store positions as waypoints, you should note the waypoint number in your notebook while in the field, and transcribe the coordinates of each location from the GPS as soon as practicable. Some models of GPS can also be connected to a computer for easy download of waypoint data.

3.2 Labelling Adequate sample description and labelling are extremely important in sampling (see Figure 3.6). Complete the labels at the sampling site and record details in your notebook. To guard against possible confusion between samples, each sample should be given a unique number. This number can be made up of parts containing codes for different pieces of information, if required. However, the label must include the following information:

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• sample location

• sample number/label

• sampler ID

• date.

Figure 3.6 Typical label on sample bottle or jar

3.3 Sample containers and preservation methods Ideally, analysis of samples should be performed in situ or at least on site. However, as this is usually not practicable, it is essential that you follow correct procedures for collection, preservation and transport of samples to a laboratory for analysis.

3.3.1 Sample containers This section applies to samples of wastes, waters and sediments. Handling and packaging requirements for animal and plant samples are discussed in Sections 3.5.4 and 3.5.5.

For samples of waters, wastes and bottom sediments, each sample should be collected and stored in a container appropriate for the quality characteristics of interest. However, if when investigating a pollution incident there is only one chance of taking the sample, and the specified type of sample container is not available, it should not be assumed that sampling is not worthwhile.

Appropriate containers and preservation methods are necessary to avoid risks of contamination of the sample and/or losses of analytes of interest during storage and transit prior to analysis. Details of sample containers and preservation are given in Appendix C8. These are based on Australian Standards AS/NZS 5667:1998 and AS 2031:2001 and overseas standards, with updating and other changes on the advice of Queensland Health Forensic and Scientific Services (QHFSS).The information in Appendix C8 is intended as a field reference with specific directions that can be followed without need for detailed knowledge of analytical procedures.

It is important that you follow these specifications exactly. If this is not possible, ensure you make a written record of what methodology you adopted.

The requirements listed in Appendix C8 include:

• the type of material/s suitable to contain the sample (container body and cap)

• the suitable method/s of pre-cleaning sampling containers

• preservation procedures

• maximum holding times

• comments on sampling procedures.

Containers should preferably be supplied by the laboratory, prepared as described in this manual and ready for use. Each container should already have a waterproof label attached with spaces for the user to fill in the

appropriate details for the sample.

Prepared containers should preferably be supplied in sealed plastic bags with some type of tamper evident closure such as a suitable seal incorporating the laboratory’s name (see Figure 3.7). All containers must be subjected to a quality control program to ensure their integrity with respect to the parameters being analysed.

Figure 3.7 Sample bottles security sealed in plastic

Once samples are taken, some of their quality characteristics can change naturally. To keep these changes to a practical minimum, for some analytes of interest specific chemicals such as acids are added as preservatives (see section 3.3.2 and Appendix C8).

If using an intermediate container, you should, ideally, have it prepared in the same way as the final one (e.g. acid-washed). If that is not practicable, and if you consider it essential to collect a sample there and then, at least ensure that the container is visibly clean, and report its preparation and use to the laboratory.

3.3.2 Preservation and storage For some quality characteristics listed in Appendix C8 there is a choice of preservation method. This is shown by a solid horizontal line separating the preservation details in the respective columns within the particular entry. A note below the name of the quality characteristic alerts you to this fact.

The amount of each preservative to be added is stated in one of these ways:

• until the sample pH reaches a stated range or

• as an amount relative to the volume of sample; this amount is stated by mass (if a solid) or by volume (if a liquid) or

• as one or more multiples of a fixed amount relative to the volume of sample, until an observable condition is met. For example, the system for preservation for metals analysis incorporated in Appendix C8 was designed to make the added preservative approximately one per cent by volume in the sample given to the laboratory.

Preservatives should preferably be supplied in small vials sealed in plastic bags, as described above for sample containers. The vials (e.g. Figure 3.8) should be labelled with the following information:

• preservative type and quantity

• preservative expiry date

• batch number

• hazard warnings if necessary.

Sometimes sample containers are supplied with preservatives in situ. This may be in the form of a small volume of liquid or crystals inside the empty container (for example, sodium thiosulphate crystals inside sterile jars intended for taking water samples for bacteriological testing). Check the supplier’s labelling in respect of included preservatives. Should included preservatives be present, then care is needed to prevent loss of preservative when filling the container, for example, avoiding overfilling or spillage.

If your sample volume is different from the stated ‘typical volume’, you must ensure that the proportion of preservative to sample remains at the intended ratio or greater.

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For characteristics where the Appendix C8 tables show ‘Refrigerate’ or ‘Freeze’ in the column headed ‘Storage conditions’, the appropriate temperature range is shown in the table heading. The sample should be cooled to this range as rapidly as is reasonably practicable and kept within that range until analysis commences (Note: AS/NZS 2031:2001 indicates 2–10°C for refrigeration).

Figure 3.8 Example of preservative container for preservation of samples

3.4 Preventing contamination Avoiding sample contamination is an important aspect of sampling. There are always potential sources of contamination, and the aim should be to keep the risk of contamination to a practical minimum, consistent with the types of analytical tests required. Possible sources of contamination include:

• Sunscreens (zinc oxide) or insect repellents (organic chemicals) on skin could contaminate a sample if transferred by some unintentional means to the material sampled.

• Where samples are being taken for both metals and nutrients, there is risk that nitric acid (used as preservative for the metal samples) could contaminate the nutrient samples. Precautions could include keeping the acid container closed while collecting nutrient samples, or having a different person collect each group of samples. More information on sampling for nutrients is given in Wruck and Ferris (1997) and QHSS (1998).

• Residual sample material from previous tests could give incorrect readings when measurements are being made with field instruments. Special attention should be given to probe and test kit item rinsing after each field measurement in order to prevent future contamination. See section 3.1.1 concerning intermediate containers.

• Avoid smoking and wear Nitrile gloves at all times.

• Corrosion and oxidisation of metal components in probe cathodes, electrodes and membranes could contaminate a sample and yield inaccurate readings.

• Note that some container caps have inserts; never touch the inserts with the skin or remove them from the caps.

• Cover work spaces used for sample handling (e.g. vehicle tray or tailgate) with new alfoil or plastic to provide a clean working surface. Replace it after driving to a new site.

3.5 Collecting samples Figure 3.9 summarises the steps in the general procedure for samples of wastewaters and surface waters.

Figure 3.9 Steps in the sampling procedure

Further detailed directions and tips for these steps can be found for some sample types in the relevant subsections.

3.5.1 Surface waters Using a waterproof pen, complete details on the label attached to the container, if possible before you collect the sample; details of labelling are addressed in Section 3.2. Be sure to let the ink dry completely before immersing the container. You might also use a strip of waterproof tape fastened around the container to ensure the label does not become detached (one end of the tape adhering to the other), and place the container into a plastic bag.

If practicable, collect the sample directly into the sample container, holding it either by gloved hand or by means of a sampling rod. If this is not practicable, collect the sample with a sampling beaker and transfer it promptly to the sample container, but beware of contamination (see Section 3.1.1).

If sampling from an open channel, try to take the sample in the centre of the channel, where the velocity is highest. Hold the mouth of the sampling container well above the base of the channel, to avoid disturbing and picking up any settled solids.

If the water depth permits, the mouth of the sample container should be held approximately 10 cm below the water surface. The surface layer of water often comprises a lipid-rich surface microlayer that may contain organic contaminants many times more concentrated than the water just below the surface.

Keep your hand out of the flow as much as possible to minimise risk of contaminating the sample, or causing infection or other harm to yourself. If the sample is to be analysed for substances present in ‘low’ concentrations—for example, nutrients—any contact of your skin with the sample should be avoided. If sampling from a boat in shallow waters, beware of stirred up sediment created by the turbulence of the boat’s movement. If this happens,

Record temperature in notebook and discard extra temperature check sample

Complete Notebook and Forms

26

do not collect the sample as soon as the boat arrives at the site. Wait until the sediment has settled. If sampling in deeper waters, avoid taking a sample in or near the wake. Take the sample from ahead using a pole, or intermediate container.

To avoid labels rubbing off sample bottles while stored and transported in eskies filled with ice, it is advisable to place bottles within individual zip-lock bags.

Avoid:

• scraping the walls of drains, tanks, sewers, and so on; this could dislodge adhering matter into the sample, and so make it unrepresentative

• disturbing sediment, if sampling in shallow waters; this could also make the sample unrepresentative of the water column.

Do not:

• smoke during operations

• rinse sample containers with waters or wastes being sampled

• risk loss of preservatives by overfilling sample bottles.

Should a problem arise while you are collecting a sample—for instance if your sample becomes contaminated by floating fats in a waste pond or disturbed sediments in a stream—use a fresh sample container and start again.

It is preferable to use the unique sample ID number rather than write descriptive text on labels. You must record the descriptive text in notes for future reference.

If taking microbiological samples from a body of water that has enough depth to immerse the sterile container, hold the container by the sides and keep it nearly upright as you lower it into the water, so that it fills without spilling any pre-added sodium thiosulphate preservative from the sample jar (if present). For situations where it is not practicable to immerse the sterile container in the water or wastes to be sampled, you might need additional equipment as follows:

• an intermediate container that can be dipped into the water or wastes and that can be sterilised (by flaming) in the field immediately before use—for example, a stainless steel jug

• a means of sterilising the intermediate container—for example, a portable butane or propane burner. AS 5667.1 states that methylated spirit flames should not be used as they are not hot enough and are difficult to control. NOTE: Flaming involves a risk of causing serious injury to people and damage to property. While flaming the jug, hold it by the handle with a sampling rod, piece of rag, or other suitable insulating material; if using a rag or other combustible material, take great care you do not set fire to it. Transfer the sample immediately to the sterile sample container. Turn out the flame as soon as you are finished.

If sampling temporary waters, refer to Appendix C4 for further information.

If sampling for suspended solids or nutrients refer to Appendix C10 for further information on sample collection with different types of sampling equipment.

If collecting bulk water samples for use in direct toxicity assessment (DTA) laboratory trials, refer to Appendix C5.

When completing your notebook, record the following sample details even if already included in a sampling schedule:

• name of the sampler

• date and time of sampling

• details of sample location and source

• quality characteristics to be determined

• container type (volume, washing, material etc.)

• sample volume collected

• preservation method used

• serial numbers of seals affixed to sample container

• unique sample identification number (a different number for each sample container and package)

• any other details relevant to the sampling, for example, photographs by number.

3.5.2 Groundwaters Groundwater sampling requires special equipment for sampling from a bore-hole or well, for example, a suitable bailer or pump, and a procedure to ensure sampling of fresh recharge water from the aquifer. Sometimes special precautions are needed to prevent changes in quality of the groundwater due to effects such as:

• reduced pressure when brought to the surface: this can cause gases in solution at the higher pressures underground to be evolved – in some cases, these can be toxic gases, such as hydrogen cyanide if the groundwater has been contaminated by cyanide solution

• exposure to components of the atmosphere such as oxygen; this can oxidise compounds naturally present in the reduced form (for example, ferrous ion).

Collecting groundwater samples needs specialised knowledge. If you need to take groundwater samples and have not been trained to do so, it is recommended that you refer to AS/NZS 5667 11 1998 (Water Sampling Guidelines—Part 11 Guidance on sampling groundwaters).

Sampling groundwater for microbiological examination requires the normal precautions mentioned above, plus the precautions for preventing microbial contamination of the sampling equipment and the sample applicable to surface waters sampling (section 3.5.1). If you need to collect such samples, you should obtain advice from reliable sources such as those cited above.

3.5.3 Sediments Sediment samples are typically more heterogeneous than water and wastewater samples. This means special care is needed in removing sediment samples from a stream bed. It also means that a composite sample is more likely to be appropriate than a single sample.

For detailed advice on sediment sampling, refer to AS/NZS 5667.12:1999 ‘Guidance on the Sampling of Bottom Sediments’ and Simpson et al. ‘Handbook for Sediment Quality Assessment 2005 (CSIRO)’.

General rules when sampling sediments include:

• If direct collection into a sample container is impracticable, use a suitable mechanical device such as a stainless steel grab, dredge, or corer, washed in waters at the sample site.

• If the sample is NOT to be frozen, fill the container almost to the brim. Chill samples in ice or refrigerate promptly.

• If freezing of samples IS required, fill the container to only two-thirds of capacity, including any cover water taken from the same site. Place samples promptly in dry ice or portable freezer.

• It may be necessary to sample sediments from temporary waters such as intermittent streams or ephemeral streams. In such cases you should refer to Appendix C4.3.2 for further guidance.

3.5.4 Fish and other aquatic animals Public complaints of ‘fish kills’ are a common reason for taking samples of aquatic animals. The Fish Kill Reporting and Investigation Manual (DEH, 1998) contains detailed advice on collecting these samples. The information below is a summary.

How many to collect

Usually, there will be many more fish than you need to collect for analysis or examination. In such cases, if some fish appear to be still alive (e.g. some might be moving but showing signs of lethargy or distress), choose them rather than those that already appear to be dead (such as any that do not move when touched, or any showing signs of decomposition, for example odour). Nevertheless, if only already-dead fish are available for sampling, they are suitable for chemical testing (but not histopathology).

Under the Animal Care and Protection Act 2001, approval from an Animal Ethics Committee is required for the sampling of live fish for scientific purposes. If approved, the handling of fish collections should be consistent with the guidelines set out in the Animal Care and Protection Act. Subject to approval, the euthanisation of aquatic fauna via chilling is a widely accepted technique consistent with preserving sample material and any related contaminants. A practical method of chilling for this purpose is to place the fish in a plastic bag of water and ice for 10 minutes or longer. In the case of fish from brackish or salt water habitats, the ice should first be placed into a separate plastic bag, in order to avoid rapid changes in salinity as the ice begins to melt.

The amount collected should be:

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• for physicochemical analyses at least three whole fish (of the same species) of approximately uniform size if possible to allow valid comparison between sites. If the fish are small, you might need to collect more than three to get enough material for analytical tests. Collect as many as needed to provide:

o for organic analyses such as pesticides, herbicides and PCBs—at least 250 grams (whole body mass) and/or

o for inorganic analyses such as heavy metals – at least 100 grams (whole body mass).

• Note: As you might not know which contaminants are present, it is usually best to collect enough material for both types of analyses.

• For histopathological examination:

• At least three whole fish (of the same species) of approximately uniform size if possible to allow valid comparison between sites. Note: histopathology requires live or freshly euthanased samples, or tissue samples preserved with 10 per cent formalin.

• Do not freeze samples.

See Part E of this manual for more detailed advice on taking and handling samples for veterinary examination.

Dissection of fish for samples of organs

If physicochemical analysis of fish organs (such as gills, livers) is to be performed by the laboratory, the fish should be dissected and the organs separately packaged, labelled and preserved before despatch.

Cross-contamination of samples can easily occur if dissection is done without proper care. Reference to the Fish Kill Reporting and Investigation Manual (DEH, 1998) is strongly recommended.

Packaging and preserving

Samples of aquatic animals and fish organs should be packaged according to the tests required as follows:

Physico-chemical analysis:

• Where poisoning by pesticides or organic compounds is suspected:

o wrap specimens in aluminium foil with the dull side of the foil inwards

o freeze as soon as practicable.

• For other analyses, including metals:

o place samples in polyethylene bags or wrapping

o freeze as soon as practicable.

Histopathological examination:

• If fish are dissected on site, follow directions in the Fish Kill Reporting and Investigation Manual for fixing the organs in 10 per cent formalin in plastic sample jars.

• Send to the histopathologist as soon as practicable.

See Part E of this manual for more detailed advice on taking and handling samples for veterinary examination.

3.5.5 Vegetation and algae Chemical analysis of aquatic or terrestrial organisms associated with the site of an incident can sometimes give information concerning the nature or source of a release of wastes or other source of contamination. For example:

• Plants could have accumulated a deposit or show discolouration on leaves or other parts.

• Aquatic plants can bio-accumulate metals and organics from the water column and the sediments.

• The leaves of emergent aquatic plants can bio-accumulate metals and organics from the atmosphere.

• Terrestrial plants can bio-accumulate metals and organics from the atmosphere and the soil.

Typical plant samples include leaves, bark and roots.

If vegetation surfaces (e.g. leaves) may have been contaminated by spray drift, consider wipe sampling (Section 3.5.6)

Plant samples should be packaged in a similar way to fish:

• If the sample is to be analysed for organics, wrap in aluminium foil with the dull side of the foil inwards, and chill or refrigerate.

• If the sample is to be analysed for inorganics, wrap in polyethylene bag or wrapping, and chill or refrigerate.

It may be necessary to collect samples from algal blooms for identification of the offending species. See part D of this manual for more detailed advice on taking and handling algal samples.

3.5.6 Wipe sampling of surface contaminants (also known as ‘swab sampling’) The presence of surface contamination (for example, by dusts, spray drift, or residues from containment) can be assessed by taking a swab from the surface concerned.

This is a useful technique for testing for the presence of contaminants on a surface. However, although there are regulatory measures of surface contamination for some contaminants (for example, for ‘PCB-free’ materials), the use of wipe samples for a purpose other than detecting contaminant presence (for example, comparing surface contamination on a spatial basis, for checking the efficacy of decontamination, or for comparison with environmental effects guidelines) is not recommended. In the latter case, the units are incompatible (surface contamination is measured in mass per unit area, and environmental effect levels are concentrations per unit volume or mass).

Wipe sampling is applicable for assessing surface contamination both from organic contaminants such as pesticides and from inorganics such as metals. Wipe sampling is most efficient on smooth surfaces such as glass, metal, painted surfaces, and smooth vegetation surfaces such as leaves. It is less effective on surfaces that are rough and/or porous such as timber and concrete.

There is no standard method for wipe sampling, although there are published standard methods for specific tasks (for example, ASTM E1728 for sampling lead dust from surfaces).

The design of a wipe sampling protocol involves choosing a material for the swab, a solvent to wet it, and a standard area to swab. A minimum of 100 sq cm surface should be wiped. If only trace levels of contamination are anticipated, wiping a much larger surface area is advisable, for example, up to a square metre. For wiping such large areas, more than one wipe may be used, and wipes can be pooled for analysis.

A variety of readily available materials are suitable for use as swabs. The portions of material chosen should be uniform in all respects and free of measurable contamination in respect of analytes of interest. The latter should be checked by analysing blank swabs. Examples of suitable swab material supplied in uniform packs are filter papers and small gauze pads such as those used for small wound dressing.

The wetting solvent of choice in most cases is an organic solvent such as isopropyl alcohol or hexane. Methylated spirits purchased in a glass bottle from a pharmacy can be used. Water is only appropriate for inorganic dusts. The purity of the solvent needs to be assured against the possibility that it is a source of any of the analytes of interest (for example, by testing a blank).

The sampling protocol should define a standard area for swabbing, as well as a standard method of taking the swab. For example, it might involve wetting each swab with 2 ml of alcohol, and wiping the swab across a pre-marked 10 cm x 50 cm surface from left to right until the whole surface has been covered, then wiping again from top to bottom, while applying an even pressure and holding the swab flat against the surface. An alternative to pre-marking a surface is to swab within the boundaries of a pre-cut template held against the surface (the template would need to be appropriately cleaned between sites). In the case of leaves, a uniform area could be approximated by swabbing the surfaces of a fixed number of leaves of similar size at each site.

Nitrile gloves should be worn and changed between sites, and used swabs sealed in labelled sampling containers appropriate for the storage of the analytes of interest, for example, a solvent-washed jar if pesticides are of interest.

3.6 Instrument-based field tests To obtain accurate results for some quality characteristics, measurement on-site is necessary.

Typical field equipment used for this is:

• thermometer

• pH meter

• dissolved oxygen (DO) meter

• conductivity meter

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• turbidity meter

• Secchi disc.

Some modern field instruments (multi-parameter instruments) can measure more than one quality characteristic, for example, both temperature and DO (see Figure 3.10). Note that not all field instruments give results of similar accuracy. It is important to check that the instrument used meets requirements and is calibrated (indicated by documentation of current calibration status).

Figure 3.10 Testing waters with a water quality meter

3.6.1 General guidance on taking field measurements

3.6.1.1 Background

Information in this subsection is provided to help you take water quality measurements in a scientifically valid manner so that the results will represent the field conditions fairly.

You should be familiar with the contents of the manufacturer’s manual specific to any instrument that you use, specifically in respect of calibration, use of the instrument and any limitations. If the instrument is not in calibration or is used incorrectly the data is of dubious value.

3.6.1.2 Instrument maintenance and calibration

Due to the variety of instruments currently available that perform these tests, it is not practical to provide instrument-specific advice on storage, calibration and maintenance in this manual. It is strongly recommended that instruments are stored, calibrated, maintained and used as per manufacturer’s instructions.

As a general principle, calibration should be checked and recorded before, during and at the conclusion of field use.

It is essential that unambiguous written records are kept of instrument maintenance and calibration. See Box 3.1 for guidance.

3.6.1.3 Take multiple readings

As with any other sampling, when taking instrument-based field measurements, multiple readings should be taken and recorded, and reference sites should be used to acquire background or ‘normal’ data for comparison purposes.

3.6.1.4 Keeping records of field measurements

Because the results of field measurements could be used in court actions, it is important to carry out all measurement procedures in a precise, consistent and reliable manner.

The original records made during measurements should be entered directly in a notebook, in case they are required for presentation in court.

3.6.1.5 Response to atypical or non-complying instrument readings

If readings appear atypical or non-complying (based on ‘acceptable criteria’ in the standard operating procedure for the instrument in question), the first step should be to check for equipment problems, such as a broken electrical cable or insulation, faulty probe, depleted batteries, etc. If equipment appears sound you might take extra measurements to confirm that the results are valid.

The value of calibration before and after instrument use, and the taking of measurements at reference sites are particularly important if atypical or non-complying readings are encountered at a site of interest.

3.6.1.6 Importance of general observations

Your observations during measurements can be extremely important in assessing atypical events or long-term trends, especially when investigating pollution incidents. Such observations could include:

• atypical water colour or clarity (such as greenish, muddy, pale brown and cloudy)

• odours

• wind speed and direction

• surface scum

• heavy algal or plant growths

• dead or dying vegetation in waterways or on banks

• dead or dying fish

• flotsam

• dumped material

• nearby earthworks or other construction activity

• nearby agricultural activities

• nearby industrial establishments or wastewater treatment works.

As visible conditions could be difficult to describe accurately in words, it is strongly advised to take photographs if possible (but note that some films have known colour bias). You should keep a record of the photograph number on the memory card/film to avoid any later confusion.

Box 3.1 Maintenance and calibration of water quality measuring instruments

All instruments used for taking measurements in the field (e.g. pH, temperature, conductivity, dissolved oxygen) must be maintained and calibrated to ensure reliability and credibility of the data they produce.

The aim is to ensure that each instrument is maintained in a sound operating condition, is capable of operating at acceptable performance levels, will not deteriorate from lack of required servicing, and that the credibility of the data the instrument produces can be demonstrated (for example, by the production of maintenance records and calibration logs in evidence).

Manufacturer’s recommendations should be the basis of maintenance and calibration, in terms of both methods and frequency. As a general principle, the calibration of an instrument should be checked before and at the conclusion of a sampling exercise. It may be advisable to also re-check calibration once or more during an extended period of use, and after transport between sites or other circumstances that may affect reliability. Such ‘in-field’ calibrations should be recorded in a notebook and later transcribed into the calibration log book for the instrument (see below). Variability in performance shown by such calibration checks before, during, and at the conclusion of a sampling exercise should be reported with the data.

Because ‘in-house’ calibrations rely on buffer solutions and other techniques, a ‘cross check’ calibration using a real water sample should be conducted from time to time. This is done using two or more instruments simultaneously to test a bucket of typical environmental water (e.g. from a creek). The readings of calibrated and reliably performing instruments should be consistent with each other for such a sample.

For each instrument the following procedures and documents should be established and kept up to date:

• a list of spare parts and suppliers/sources of repair (it may also be feasible for a realistic supply of user-replaceable items to be kept on hand)

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• written inspection, maintenance and calibration schedules (based on the manufacturer’s documentation and the usage pattern for the instrument)

• a log book in which is recorded inspection, maintenance and repair activities detailing dates and persons involved. This may best be documented as a ring file arranged in chronological order covering the service life of the instrument. Sample inspection/maintenance records are provided in Appendix C1

• a log book in which is recorded calibration activities detailing dates, times and persons involved. This may best be documented as a ring file arranged in chronological order covering the service life of the instrument. Sample calibration records (log sheets and charts) are provided in Appendix C1

• adequate supplies of calibration standards (if these are required) should be maintained and where applicable storage and shelf life requirements followed.

Some of the above documents may be combined, for example, the maintenance schedule could be combined with the maintenance records, and the calibration schedule with the calibration records.

It is strongly recommended that within each work unit the oversight of instrument inspection/ maintenance/calibration be tasked to a specific person/s as an unambiguously assigned responsibility. Such assigned person/s would be expected to personally perform or supervise the performance of all tasks and completion of related records according to the established schedule/s for the instruments held by the work unit.

3.6.2 Overview of field measurements

3.6.2.1 Temperature

Accurate temperature measurements are required for accurate determinations of pH, specific electrical conductance, and dissolved oxygen. You need to be aware whether the instrument you are using compensates for factors such as temperature when measuring another parameter, or whether results need to be adjusted by calculation.

Stratification is common in summer months when surface waters are much warmer than bottom waters. Accordingly, unless the water is shallow (less than 0.5 metres) and flowing, take temperature readings at different (measured) depths.

Warm water is less capable of retaining dissolved oxygen than cold water. For this reason, temperature should be measured at the same place within the stream at which dissolved oxygen is measured. This allows the correlation between the two parameters to be observed.

3.6.2.2 pH

A pH of 7 is called neutral, above 7 is basic (or alkaline), and below 7, acidic. Strong mineral acids such as concentrated phosphoric acid can have pH less than 1; strong alkalis such as caustic soda solutions can have pH approaching 14.

3.6.2.3 Dissolved oxygen (DO)

The maximum concentration of dissolved oxygen (DO) in water under ambient conditions is typically within the range of about 6–10 mg/L, depending on the atmospheric pressure and the water temperature and salinity.

Dissolved oxygen concentrations are most often reported in units of milligrams of oxygen gas (O2) dissolved in each litre of water, i.e. mg/L (the unit mg/L is equivalent to parts per million = ppm).

An alternative measure is dissolved oxygen saturation (per cent). This is the percentage of dissolved oxygen concentration present relative to what the concentration would be if water at the specified temperature and salinity was fully saturated with dissolved oxygen. Most dissolved oxygen probes compensate automatically for temperature and salinity when calculating dissolved oxygen saturation in water (check the instrument manual). Note that under natural conditions such as high algal density during sunlight, super-saturation (more than 100 per cent DO) can occur.

Considerable differences between DO concentrations at the surface and in the lower depths can result from stratification of the water column, due to temperature or salinity effects. This effect is usually most pronounced in summer months when surface waters are much warmer than bottom waters. Accordingly, unless the water is shallow (less than 0.5 metres) and flowing, take readings at different (known) depths.

3.6.2.4 Electrical conductivity (EC)

Electrical conductivity (abbreviated EC, and often simply called ‘conductivity’), the ability of water to carry an electric current, is used as an indicator of salinity and the concentration of dissolved salts in a waterbody.

The unit of measurement for conductivity is siemens (S) per unit of length. A commonly used example of this unit is microsiemens per centimetre (µS/cm). See Conductivity units and their abbreviations in Appendix C7 for further explanation.

Typical values of EC in µS/cm are:

• De-ionised water in equilibrium with the atmosphere approximately 1

• Potable waters 50–500

• Freshwater less than 1500 (varies widely between catchments)

• Seawater approximately 52 000

Note that conductivity varies with temperature, and values reported are usually those corresponding to 25OC. A difference of 5OC can alter conductivity by approximately 10 per cent. Many conductivity meters have compensation functions so that EC at 25OC can be read directly. However, if necessary, a manual correction can be made by using the formula

25) - ( 0.019 1

t25

+=

tKK

where t = water temperature OC where conductivity is measured

Kt = conductivity at temperature t OC

K25 = corrected (25OC) conductivity of the water

3.6.2.5 Relationship of EC to salinity and dissolved salts/solids

Confusion can arise from the fact that salinity is sometimes equated to one of the terms ‘total dissolved ions’ or ‘total dissolved salts’ or ‘total dissolved solids’, and that the same acronym (TDS) may be used for the last two terms. The following notes explain the distinction.

• ‘total dissolved ions’ is the sum of the ion concentrations. This sum is used in a formula to calculate ‘total dissolved salts’.

• ‘total dissolved salts’ is determined by calculation from the results of analysis for common ions (e.g. sodium, calcium, chloride).

• ‘total dissolved solids’ is determined by filtering a sample, drying at a specified temperature, and weighing the residue. It includes non-ionised species if present (e.g. sugars, other organics, colloidal particles too small to be retained by filter medium), with the result that ‘total dissolved solids’ can be greater than ‘total dissolved salts’.

For typical fresh waters, the approximate total dissolved salts can be calculated from conductivity using the formula:

TDS = 0.68 x K25

Where TDS = Total Dissolved Solids (in mg/L)

K25 = conductivity of the water at 25 OC (in mS/cm)

Note—this formula is not appropriate for water of atypical content.

Salinity in parts per thousand (g/L) can be calculated from conductivity at 25 OC using the formula:

S = a1(K25) + a2(K25) 2 + a3(K25) 3 + a4(K25) 4 + a5(K25) 5 + a6(K25) 6

Where S = salinity, g/kg, % or ppt (approx. equal to g/L)

K25 = conductivity of the water at 25 OC (in mS/cm)

a1 = 4.980 x 10-1 a2 = 9.540 x 10-3 a1 = -3.941 x 10-4

a1 = 1.092 x 10-5 a1 = -1.559 x 10-7 a1 = 8.789 x 10-10

3.6.2.6 Turbidity

The turbidity of a water body is a measure of the presence or absence of soluble, suspended and colloidal particles that hinder the transmission of natural light from the surface to the lower depths. Turbidity affects the potential rate of photosynthesis, and hence the growth of plants or algae in the water body.

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3.6.2.7 Water clarity

The clarity of a water body is an indication of the presence or absence of suspended and colloidal particles that hinder the transmission of natural light from the surface to the lower depths. Clarity affects the potential rate of photosynthesis at any given depth, and hence the growth of green plants or algae in the water body.

This method is based on the common experience that the deeper a submerged object is, the less easy it is to see from the water surface, and the more cloudy the water, the less easy to see at a fixed depth. It entails the use of a Secchi disk (see Figure 3.11 and Figure 3.12) and is a relatively simple and quick way to obtain a measure of clarity, without the need to take samples and analyse them for turbidity or suspended solids. The Secchi disk also has the advantage of integrating turbidity over depth (where variable turbidity layers are present).

Figure 3.11 Testing clarity of water using a Secchi disk A Secchi disk is not appropriate for use in shallow waters where the disk can still be seen when resting on the substrate. In such cases you should use a turbidity meter or take water samples for laboratory analysis.

NOTE: The observer’s visual acuity will affect the perception of the disc. It should be observed with ‘corrected vision’ (spectacles or contact lenses if these are normally worn) and not be performed on the side of the boat casting a shadow. Tinted lenses or sunglasses should not be worn, as they could affect the depth where the disc is recognised.

3.6.2.8 Chlorine (free, combined, and total residual)

Chlorine is widely used for disinfection of public water supplies, swimming pools, and treated wastewaters. Although the role of chlorine addition in water treatment is to disinfect (kill pathogens), some of the added chlorine can be consumed in reactions with oxidisable substances present in the water including ammonia, nitrite, and organic matter. ’Chloramines‘ are a common product of such reactions, which also deplete the chlorine available to kill pathogens. The reaction products are generally less effective disinfectants than chlorine, but are more persistent.

Chlorination treatment of water or wastewater usually involves the addition of a measured dose of one or more of chlorine gas (Cl2), hypochlorite ion (OCl-), and hypochlorous acid (HOCl). The addition of chlorine gas alone results in a mix of all three in proportions dependent on factors such as pH and temperature.

The term ’free chlorine‘ is used to refer to the total concentration present of dissolved chlorine gas, hypochlorite ion, and hypochlorous acid, each of which has good disinfection capability.

The term ’combined chlorine‘ is used to refer to the chloramines, and the term ’total residual chlorine‘ is the sum of ’free chlorine‘ and ’combined chlorine‘. These are the terms used in the Australian Guidelines for Water Quality Monitoring and Reporting (ANZECC/ARMCANZ 2000). Other terms for these mixtures of chlorine-containing substances may be encountered, for example some environmental authorities (EAs) include quality specifications for ’free chlorine residual‘ in an effluent. The term ‘residual’ refers to chlorine (as the total concentration of dissolved chlorine gas, hypochlorite ion, and hypochlorous acid) remaining after added chlorine has reacted with wastewater constituents. Thus ’free chlorine residual‘ is equivalent to ’free chlorine’.

The quality specification for chlorine (free, combined, or total) in an EA is usually the concentration measured at the outlet of a ‘detention tank’ designed to provide contact between the chlorine and the wastewater for a stated period to enable disinfection to occur prior to release.

Because the levels of ’free chlorine‘ relative to ’combined chlorine‘ can change over a short period of time, it is usual to measure chlorine in situ using a test kit or probe attached to a water quality meter. There are more sophisticated and accurate laboratory based methods available but these are generally not practical for field use.

Test kits commonly used for in situ chlorine testing are based on a colorimetric system, involving the addition of a chemical (DPD, typically supplied in the form of tablets in a sealed containment) to a fixed volume of sample water and measuring the intensity of pink colour produced by the reaction of the DPD with chlorine present in the water. The methods used to measure the colour intensity vary between kit types. Simple kits involve comparison by eye of the colour intensity with a calibrated chart or filter. More sophisticated (and accurate) kits measure colour intensity digitally. The results from DPD-based test kits may be adversely influenced by colours and interferences from chemicals present in the waters being tested (see section 3.6.3).

If using a DPD kit, it is essential to ensure that the DPD tablets are fresh (hard), that the seal is intact, and within the expiry date. The kit directions for use must be followed exactly, and cells and colour filters etc. kept scrupulously clean. Residues from previous tests can give false readings for subsequent tests.

3.6.3 Test kits A range of commercial field kits are available for rapid testing of water quality. These include tests for analytes such as pH, metals and nutrients (among others). Test kits are a substitute (surrogate) for a range of instrumental techniques of which probes are just one. These kits can provide valuable on-the-spot information, although caution is advised in their use. It should not be assumed that a test kit will perform as specified by the manufacturer.

There are many factors which influence any instrument’s performance. Test kits are often subject to a range of limitations and interferences depending on the type and nature of water being analysed (sample matrix). It is imperative test kit performances are initially verified to ensure they are fit for their intended purpose. This can be achieved by undertaking validation experiments covering the analytical range and the matrices being investigated. Once validated, acceptance criteria can be ascribed to the various tests. These acceptance criteria may or may not align with the manufacture’s specifications. Some level of quality control, such as use of check standards, must be utilised and monitored each and every time they are used.

It is important to remember the following points:

• Test kits are utilised in field environments and can be subjected to extreme physical conditions which will impact on kit performance. How do you know the test kit is functioning adequately? Has its performance characteristics changed since the last time it was used?

• Most kits have expiry dates for reagents that should be checked and adhered to.

• Test kits will require storage in an appropriate fashion (e.g. within certain temperature ranges; out of direct sunlight).

• Test kits often have varying levels of accuracy (e.g. X ± y units). Will this accuracy suit your application? (e.g. will the accuracy of this kit allow you to differentiate between sites?).

• Test kits often have varying detection limits (i.e. only detect down to a certain concentration). If comparing to guidelines/standards/benchmarks, does the kit report within the range of these documents?

• Test kits can behave differently in fresh, waste and salt waters.

• Water containing particulate or suspended matter (even at small concentrations) can severely impact on the test kit producing a reliable result. For these types of samples, they should be filtered appropriately before using the test kit.

• Different brands of kits may vary in all of the above points.

N.B. It is NOT advisable to rely on test kits for data that may be required for legal purposes.

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3.7 Sample security and transport

Figure 3.13 Overview of the components of a sample security and transport system

3.7.1 Securing your samples It must be possible to demonstrate that there was minimal risk of the sample having been interfered with between the time of sampling and the time of analysis. This requires a well designed system for security of the samples, including precautions to make any such interference evident upon receipt by the analyst. The system described in this manual is summarised in Box 3.2 and should be adequate for most purposes.

3.7.1.1 Sample seals or evidence bags

Sample seals and evidence bags can be obtained from a range of commercial suppliers.

Typically seals are specially printed self-adhesive ‘security’ labels, designed to be affixed across the body and cap of the sample container. Each seal is made of a ‘self-destruct’ material so that any attempt to remove it will result in its disintegration so it cannot be re-affixed in its original condition. Each should have a unique seal number. Uniquely numbered and tamper-proof tough-plastic ‘evidence bags’ are commercially available in a range of sizes.

When you are issued with seals and evidence bags, you are responsible for keeping them in a secure place. At any time you must be able to account for the numbers received, the numbers used, and the numbers still in your possession. Include the seal/evidence bag number and the sample identification number when recording details of samples. Refer to Figure 3.14 for examples of security labels and seals.

Sample Security and Transport Sample Security and Transport

Securing your samplesSecuring your samples Transporting your samplesTransporting your samples

Are your samples tamper-evident?Are your samples tamper-evident?

Are you delivering the samples?Are you delivering the samples? Is someone elsedelivering your samples?

Is someone elsedelivering your samples?

Documentation Chain of Custody etc

Documentation Chain of Custody etcSample ReceiptsSample Receipts

Samples packaged properlyto avoid damage?

Samples packaged properlyto avoid damage?

Have your samples arrived?Have your samples arrived?

Have your samples been analysedwithin maximum holding time?

Have your samples been analysedwithin maximum holding time?

Complete analysis requestComplete analysis request

Contact LaboratoryContact Laboratory

Figure 3.14 Examples of security labels and seals

3.7.1.2 Locked carrier boxes

One way to hinder unauthorised access to samples is to use a system of insulated carrier boxes fitted with locks that can be opened only by:

• an appropriate staff member of your organisation having authority to do so

• the analyst or other laboratory staff member having similar authority.

If using such a system you should be able to testify in court that the keys were kept in secure places.

A person wishing to interfere with samples might try to remove the lock and later replace it without leaving evidence of their actions, such as cuts or holes in the box or the lock assembly. Each lock should be fitted to make this as difficult as practicable. Any attempt at access should leave evidence that laboratory staff will notice.

Each lock should preferably be a part of the body of the carrier box, or a padlock that fastens a hasp and staple assembly permanently fitted to the body. The two parts of the assembly would need to be fastened by, for example, suitable rivets, rather than screws that could be removed and replaced without leaving evidence of the fact. Padlocks should preferably be case hardened to resist cutting by a hacksaw.

3.7.2 Transporting your samples

3.7.2.1 General

After collecting the samples, you must have them transported to the appropriate laboratories, ensuring that the integrity of the samples is maintained. You can do this by:

• delivering them personally

• having a work colleague deliver them

• sending them by commercial carrier (such as road transport, air cargo)

• other reliable means.

Whatever means you use, the following points are important:

• Sample containers and packages need to be packed in sample carrier boxes to minimise the risk of breakage, leakage or spillage during transport.

• Sample carrier boxes must be handled and stored so as to protect the samples.

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• Appropriate security measures must be in place.

• Documentation must accompany the samples.

If samples are transported by an air carrier, they are subject to the International Air Transport Association (IATA) Dangerous Goods Regulations. These regulations are updated annually, and failure to comply with them can lead to prosecution of the consignor. It is wise therefore to consult the airline company or the Civil Aviation Safety Authority (CASA) before sending samples, to check that the sample packaging (including the carrier box) and the labelling of the carrier box meet the requirements.

If you give samples to someone else for transport, such as a commercial carrier, it may be wise to contact them at reasonable intervals afterwards to check on the actual location of the samples, or whether they were actually despatched (by road or air, for example) at the time necessary to ensure delivery within the recommended maximum holding time.

Samples should be analysed within the maximum holding times specified; otherwise the analyst must report on the probable effect of the delay.

3.7.2.2 Transport precautions

Take the following precautions when preparing samples for transport:

• Re-check caps on containers to make sure they are tight and properly sealed, so that if any should fall over they will not leak. If necessary, tighten cap, re-tape and re-seal.

• Separate glass bottles from each other to prevent physical contact, which could cause breakage.

• If you use a transportable freezer to freeze samples in the field, but need to transfer them to an insulated sample carrier for transport (for example, by a freight contractor), add dry ice to the sample carrier to keep the samples frozen during transport. Contact the freight company (air or land) to discuss refrigeration options for your samples if required.

• Ensure that the following legends are plainly readable on the sample carrier boxes: o ‘FRAGILE’

o ‘HANDLE WITH CARE’

o ‘THIS SIDE UP’

• To minimise thawing of ice or dry ice, protect samples from heat; for example, do not leave sample carrier boxes in the sun for long periods.

• Carrier boxes containing dry ice must not be hermetically sealed. Venting must be provided to release the carbon dioxide gas generated.

• Make every effort to minimise delays in transporting samples.

• Complete chain of custody documentation (Appendix C1).

• Where samples are carried by air, the carrier box must be leak-proof, and must have labels and complete documentation as required under the IATA regulations.

• Ensure that the laboratory will receive the completed analysis request either with the samples or earlier (for example, by fax or email).

BOX 3.2 Summary of sample security system

What the sampler does:

Collect sample.

Place sample in labelled container and put cap on container.

Fasten security seal across lid and container or place sample in security bag.

Put sample in sample carrier box (cooler or other type) or freezer.

Secure carrier box or freezer by:

• a lock

or

• a padlocked chain, fastened around the body so that the lid cannot be opened without unlocking the padlock or cutting through a chain link or padlock.

NOTE: the chain links should be of welded construction (NOT able to be opened and closed using pliers) and the padlock body should be a solid casting (NOT laminated from sheet metal). Padlock and chain should preferably be case-hardened.

Give sample carrier box or freezer to transport carrier with chain of custody sheet (Appendix C1) attached and get receipt (on consignment documents).

Fax or email the completed analysis request to laboratory.

What the laboratory does:

Receive sample carrier box or freezer from transport carrier, complete chain of custody sheet and give receipt (on consignment documents).

Check:

• locks and chain are intact

• there are any other signs of tampering with the carrier box or freezer.

Unlock sample carrier box or freezer and remove sample container/s.

Check:

• all samples stated in the completed analysis request are present

• security seals are intact

• there are any other signs of tampering with the samples.

If there is reason to suspect tampering, contact sampler immediately and give details.

Determine whether samples should be analysed or not.

Record results of checks and any action taken because of them.

3.7.2.3 Addresses for despatch

It is suggested you keep a list of names, addresses and contact numbers (phone, fax and email) of appropriate staff at the laboratories that your organisation employs to perform analyses, for ready reference in the field. Appendix C6 contains space for you to enter these details.

3.7.2.4 Delivery of samples

The receiving laboratory should be advised in advance that samples are to be dispatched. If you notify the staff by mail or facsimile, you should get an acknowledgement. Only in exceptional circumstances should you send samples without this prior notification.

Samples delivered to the laboratory must be handed to a supervisor or other appropriate responsible staff member. This person should acknowledge receipt of the samples by signing the consignment documents accompanying each sample carrier box, chain of custody documentation or other appropriate form of receipt.

On receiving samples, the analyst must first check the contents against the analysis request, noting:

• the time and date when the samples were received

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• whether the carrier box was locked when received

• any irregularities such as breakages or missing samples

• the condition of the samples—whether they are frozen, and if not, their temperature

• the condition of the seals and tape on sample containers and packages.

The purpose of this is to check on how well the samples were preserved, and whether there are any signs of tampering. The analyst must contact the sampler promptly if there are any signs of tampering or other irregularity, such as missing samples, broken or damaged containers, thawed samples that should have still been frozen, or samples at room temperature instead of chilled. In such a case, you should discuss with the analyst and decide whether analysis should proceed or whether you will re-sample. Keep a record of the discussion.

The analyst’s report should include details of the condition of the samples on receipt. If any irregularity was discovered on receiving the samples, the analyst must also give details of this.

3.8 Laboratory analysis

Figure 3.15 Essential laboratory analysis components for incorporation into a sampling protocol

3.8.1 Selection of analytical methods Unless otherwise prescribed, samples should be analysed, as a minimum, in accordance with a method specified in one of the following reference texts (see References section):

• APHA AWWA Standard Methods for the Examination of Water and Wastewater (current version)

• USEPA (current version)

• ASTM, Annual Book of ASTM Standards (current version)

• relevant Australian Standards published by Standards Australia, as amended from time to time

• relevant ISO Standard (current version).

The method of analysis should be chosen by the analyst so as to be appropriate to the type of sample and to the expected concentration range of the constituent to be measured. The methods used by a laboratory should be verified or validated (for example, by using standard reference materials or standard addition techniques) and

Laboratory AnalysisLaboratory Analysis

Appropriate analytical method?Appropriate analytical method?

Appropriate detection limit?Appropriate detection limit?

Appropriate precision?Appropriate precision?

Appropriate QA/QC?Appropriate QA/QC?

preferably be a method accredited by the National Association of Testing Authorities (NATA) or shown to be at least equivalent. Alternative methods to those described in the reference texts can be used provided that the analyst can validate the alternative method and prove that the results so obtained are equivalent to the results obtained using the prescribed method within the limits of the accuracy stated for the prescribed method.

3.9 Data analysis and interpretation

Figure 3.16 Essential data analysis and interpretation components for incorporation into a sampling protocol

3.9.1 Sources of reference values Once you have results from field measurements and laboratory analyses, you will need to be able to draw some conclusions based on sound scientific reasoning. The following resources act as a guide to the levels and indicators of water quality that your results should be compared against.

Reference documents useful for benchmarking environmental water and sediment quality data:

• ANZECC/ARMCANZ (2000) Australian and New Zealand Guidelines for Fresh and Marine Water Quality

• Environmental Values and Water Quality Objectives scheduled under the EPP Water, available through the department’s website at Schedule 1 of EPP Water (including plans)

• NHMRC (2004) Australian Drinking Water Guidelines

Data Analysis & InterpretationData Analysis & Interpretation

Check data integrity?Check data integrity?

Analyse dataStatistics – means, variance etc

Analyse dataStatistics – means, variance etc

Interpret dataInterpret data

Suffice study objectives?Suffice study objectives?

YesYes NoNo

Prepare dataPrepare data

Analyse changesin time and spaceAnalyse changesin time and space

Explore relationships betweenmeasurement parameters

Explore relationships betweenmeasurement parameters

Compare test site data toguidelines and/or objectivesCompare test site data to

guidelines and/or objectives

ReportReport

Refine objectives and/or

collect new data

Refine objectives and/or

collect new data

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• NHMRC (2008) Guidelines for Managing Risks in Recreational Water

• Queensland Water Quality Guidelines

• Queensland Environmental Protection (Water) Policy 2009.

3.10 Data custodianship, management, and submission for regulatory purposes Reliable and defined custodianship and data management is essential to ensure data is collected, maintained and used appropriately.

Data typically relates to measurements or statistics from measuring devices or observations, usually presented in a numerical or structured format. A collection of data may be referred to as a dataset, usually held electronically in databases (either as a collection of related data stored together in one or more computerised files, or an electronic repository of information accessible via a query language interface).

Generic classification of data includes time series data, spatial data and metadata:

• Time series data are a set of observations, results, or other data obtained over a period of time, often at regular intervals.

• Spatial data are data that refer to specific geographic areas. Records would generally include a geographic reference, for example, map references, latitude and longitude references, river catchment areas, local government areas, or others.

• Metadata is information which describes the content, quality, condition, and other appropriate characteristics of the data. The term information is a broader term often used to describe any data that is processed, organised or classified into categories, images, graphs, etc. for a designated purpose.

Good custodianship of data is needed to provide accountability for data sets and give users confidence with the level of integrity, timeliness, precision and completeness of data sets. All data, whether they are generated by government or obtained from an external source, must be managed by a custodian. A data custodian can be defined as a person or organisation that is responsibility for ensuring specific data is collected, maintained and made available according to standards and policies or other licences, agreements or specifications.

Custodians are responsible for ensuring that the following minimum standards are applied to each dataset:

• documentation of the methods and process for data collection

• ensuring data is fully validated and quality assured with sufficient detailed metadata to enable its use by third parties without referring to the originator of the data

• the dataset must be recorded to enable the ownership, access constraints and licence conditions to be determined

• a contact for allowing the release or use of data by other parties.

There are a number of situations where individuals and organisations may need to submit water monitoring or other related data to the Queensland Government for the purpose of regulatory decision making. This could include but is not limited to purposes such as environmental impact statements, development approvals, environmental management plans and environmental evaluations.

In many cases, water monitoring data is required to be submitted electronically. The provision of correct and accurate data is the sole responsibility of the submitter and water monitoring data should be collected in accordance with this document and other relevant standards, guidelines and policies. The Queensland Government will not be held responsible for submission of incorrect data. The Queensland Government also reserves the right to use submitted monitoring data for any purpose it sees fit including supply of data to a third party.

Refer to Appendix C1 for further guidance and checklists of what information needs to be included when submitting electronic water monitoring data.

Part C Appendixes

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Appendix C1 Forms Contact the department for a copy of the forms mentioned in the manual—these include:

• Sampling Schedule for Waters, Waste, Sediments

• Checklist of Equipment and Materials

• Analysis Request

• Chain of Custody Data Sheet

• Examples of Field Instrument Maintenance and Calibration Record Sheets.

Appendix C2 Methods for overcoming limit of detection problems: in situ extractions and the use of passive samplers The standard grab sample water bottle has a volume of 1 litre or less. However, this is an insufficient volume for the analysis of many contaminants at the sub-microgram per litre concentrations necessary for assessing compliance with water quality guidelines. In principle the level of detection could be improved by collecting a larger sample (e.g. a 10 litre volume could provide an order of magnitude improvement over a 1 litre volume) but this presents many practical difficulties of preservation, transportation, storage and extraction.

More practical options available include in situ extraction of the contaminants of interest, or the use of passive sampling devices.

In addition to resolving level of detection limitations, such techniques (especially passive sampling) provide a practical means of time-integrated sampling whereby contaminants can be monitored on a continuous basis for extended periods (up to several weeks at a time).

These sampling techniques provide an additional set of tools useful for modern monitoring programs.

C2.1 In situ extraction One such system, solid phase extraction (SPE), makes use of the tendency of contaminants with low water solubility to bind to specific solid materials.

The main advantage of in situ concentration is that large amounts of water can be extracted in the field and need not be transported. In addition, because these systems can be deployed for long periods, they can provide a relatively inexpensive method for integration of contaminants over time. The basic elements for such a system (piping, filter/s, sorbent column/s, pump, power supply, flow meter, control system) can be assembled readily by a good laboratory workshop. Such a system is illustrated in Figure C1.

Figure C1 An in situ sampling system pumps a measured volume of water through a sorbent cartridge

Due to the risk of contamination during field extraction of water samples using SPE, it is highly desirable to avoid contact between the water being sampled and the mechanical components of the pump prior to the water reaching the solid phase. Generally, this necessitates plumbing the pump so that it sucks rather than drives water through the extracting media. In practice this requires that the sampling device be situated as close as practical to the level of the water source because suction is limited to one atmosphere of negative pressure, and in practice, due to inherent resistances in the system (piping, filter/s and extracting media), 1–2 m altitude difference is the maximum achievable.

The risk of introducing contaminants from tubing and other components of an in situ sampler is similar to that inherent in automatic samplers (see section 3.1.2). Other requirements of QA/QC also apply such as the wearing of

46

gloves to avoid contamination of sampler components that come in contact with the sample.

C2.2 Passive sampling devices In principle the use of a passive sampling device involves the deployment of a chemical-absorbing material in the water column (or sediments). After a period of exposure, the absorbent material is retrieved and the accumulated chemicals analysed. Passive samplers can be used to detect contaminants and also to estimate average exposure concentrations during the period of deployment. The latter can be calculated from the duration of exposure, the measured concentration of contaminant accumulated, and the absorption rate of the material used in the sampler.

The basic components of a passive sampling device are an accumulating medium, a membrane to control the rate of uptake, and a mounting structure to contain and protect these components but at the same time expose them to the water being sampled.

Passive sampling devices can be deployed in the field in a variety of situations—hung from floats, suspended from jetties, fastened to stakes inserted in a stream bottom, or anchored to the bottom but held up into the water column by a float.

The normal requirements of QA/QC apply when sampling with passive sampling devices. These include the use of field blanks, replicate samplers, and precautions such as the wearing of gloves to avoid contamination of the sampler or its housing during handling.

Time-integrating passive sampling techniques have become widely used in the last decade. In particular, the use of performance reference compounds introduced into the sampler to enable adjustment of field data from the samplers using kinetic data from the laboratory has increased user confidence in these sampling techniques.

A variety of accumulating media for passive sampling are available, allowing a choice based on the chemical properties of the target contaminants. Examples of common types are as follows.

C2.2.1 Passive sampling for organic pollutants

C2.2.1.1 Semi-permeable membrane device

The most widely used passive sampler design for non-polar chemicals in water is the semi-permeable membrane device (SPMD) shown in Figure C2. It consists of a length of sealed lay-flat PE tubing (the membrane) containing a small volume of triolein (the absorbent phase) woven around a stainless steel frame. The device is then inserted into a perforated stainless steel shroud for protection from mechanical damage during deployment.

Trace levels of contaminants that cannot be detected in conventional water samples are often concentrated to detectable levels by SPMDs or similar devices placed in water for a controlled exposure period.

An alternative to the triolein-based SPMD described above is to use a strip of silicon rubber such as polydimethylsiloxane (PDMS) placed inside the deployment shroud as the absorbent material.

Figure C2 Housing and components of an SPMD passive sampler. The protective cage covers the absorbent strip during deployment

C2.2.1.2 Chemcatcher

This device is a very robust passive sampling device that employs the C18 Empore disk as the absorbent media,

combined in most cases with a membrane that allows diffusion of polar chemicals. Depending on the polarity range of the analytes of interest, a range of devices incorporating appropriate membranes and solid phase absorbent media have been developed. One of these devices based on the Empore disk is illustrated in Figure C3.

4

7 5

6

8

10 11

1

Figure C3 Housing and components of a Chemcatcher passive sampling device

The sampler consists of three interlocking sections (2, 3, 9) manufactured from polytetrafluoroethylene (PTFE) that screw together during deployment to form water-tight seals (4, 10).

Integral to the device is a 50 mm rigid PTFE disk (7) designed to support both the absorbent material (Empore) (5) and the diffusion-limiting membrane (6).

On the reverse is a lug (1) for attaching the device during deployment.

The outer surface of the diffusion-limiting membrane is protected from mechanical damage during deployment by a mesh (8) of either stainless steel for organic analytes or nylon for inorganic analytes. This mesh is held in place during deployment by a removable PTFE ring (9).

A PTFE screw cap (11) replaces the ring (9) during transport to and from the deployment site.

C2.2.2 Passive sampling for metals

The diffusive gradient in thin film (DGT) device employs a resin gel as the accumulating absorbent media, overlaid by a filter (to exclude particulates) and a diffusive hydrogel to maintain a concentration gradient, with the whole loaded into a cylindrical plastic housing (see Figure C4). A series of different gels have been developed to sample a range of metals (both labile and organic species), phosphorus, sulphides and radioactive caesium, and are available in a range of thicknesses. Parallel deployment of two DGT units assembled with different diffusive gel thicknesses allows accurate measurement under low flow conditions.

An important advantage of using DGT to measure metals in saline or marine waters is that the gels do not accumulate the major ions that often cause interferences in the analysis of metals in grab samples of water. The devices sample satisfactorily over a range of pH, with the range limits varying between metals (e.g. down to pH 2 for Cu, but only 4.5 for Cd) and over the range pH 2 to 9 for phosphate.

A variant of the device, known as DET (diffusive equilibrium in thin films) can be deployed to perform relatively rapid (within a day) response times and has the ability to measure at high spatial resolution. The DET comprises a single relatively thick sheet of gel (typically 0.8 mm) supported in a holder. Solutes in the surrounding water diffuse into the gel until concentrations in gel and water are equal.

DGT and DET devices, for both of which the University of Lancaster (UK) holds a worldwide patent, are available commercially, either pre-assembled or in kit form (gel disks and strips for local assembly). For details of supply visit the DGT for measurements in waters, soils and sediments website at <www.dgtresearch.com>.

Exposure to water column


1 7 2

3

5 6

9

48

Figure C4 Housing and components of a DGT passive sampling device


cross-section

enlarged

Appendix C3 Flow measurement The most relevant Australian standard for flow measurement of water bodies is AS 3778, parts 1 to 6—Measurement of water flow in open channels. This standard is also relevant to some types of flow measurement for point source discharges. The standard describes comprehensively the range of available methods for measurement. For wastewater discharges, AS 2360—Measurement of fluid flow in closed conduits may also be relevant. Important aspects of these two standards are discussed briefly below.

Methods for measuring water flow in open channels are mainly divided into velocity-area method, measuring structures, dilution method, slope-area method and cubature method. AS 3778.21 (2001) provides guidance on selecting the appropriate methods and explains uncertainties associated with each method.

Velocity-area methods are described in detail in AS 3778 (Part 3). The velocity-area methods are commonly used for estuaries and freshwater streams where no physical measuring structures exist. Some of the methods can also be adopted for measuring wastewater flow in open channels. Most common methods for instantaneous measurements include using velocity meters (stationary or on a boat) and timing a float travelling over a known distance (freshwater only). Continuous measurements can be made using ultrasonic (velocity of sound in water) meters and electromagnetic induction in buried coils.

Weirs, flumes or other physical structure work on the principle that volumetric flow rate is related to a known cross-sectional area or height before some fall. This type of method is often used for freshwater gauging stations and can be used for measuring wastewater flow in an open channel. Some type of water measuring device that needs to be calibrated is usually required for continuous measurement.

The dilution method involves the use of a tracer and sampling of waters to measure the concentration of the tracer where it has uniformly mixed across the cross-section of the waterway. This method of flow measurement is only applicable to freshwater streams. However, tracers are commonly used in estuarine and marine waters to study the dispersion of a wastewater release and to calibrate the hydrodynamics of water quality models.

Methods for measuring flow in closed conduits such as pipes are mainly divided into pressure differential methods, mass methods and volumetric methods. Calibration is essential for this method and strongly linked to the uncertainty of the measurement (see AS 2360.7.1/AS 2360.7.2).

Pressure differential methods are commonly used for measuring flow into plants such as sewage treatment plants and may be used to infer the discharge quantities where flow losses are minimal. Confidence in the flow measurement is strongly dependent on appropriate calibration and should be undertaken as per AS 2360.7.1.

The main types of volumetric methods include static and dynamic gauges, diverters and flow stabilisers. The most common volumetric method used for wastewater discharges is a flow stabiliser where a device such as a pump is used to ensure a stable flow-rate is supplied. If the flow is not continuous, the duration for which the water is flowing can be recorded to calculate total flow for a known period.

Regardless of the method used, it is important that overall uncertainty in the flow measurement is known or calculated. Refer to AS 3778.2.4 for estimation of uncertainty of a flow-rate measurement in open channels and AS 2360.7.1 and AS 2360.7.2 for assessment of uncertainty in calibration and use of flow measurement devices for closed conduits and the specific subsections of AS 2360 for the method adopted.

Appendix C4 Sampling water quality in temporary waters Temporary waters predominantly occur within arid and semi-arid regions in Australia and are by nature highly variable systems. Temporary waters include:

• intermittent waters: areas that are predictably inundated each year, but whose duration of water retention may vary

• ephemeral waters: areas that only contain water after irregular rainfall or flow events.

Before any sampling of temporary waters begins, thorough consideration should be given to all the environmental and discharge factors presented below. Regional departmental officers are advised to consult with the Queensland Department of Environment and Heritage Protection Environmental Sciences division while others are urged to seek advice from environmental consultants experienced in developing sampling programs for temporary waters.

C4.1 Sampling the receiving environment Due to the highly variable nature of receiving environments in arid and semi-arid areas, there is no standard set of sampling techniques. Choosing the correct approach will depend on a variety of factors, which include the:

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• stage of the hydrocycle (flowing, pooling or dry)

• amount of vegetation in the catchment

• underlying geology of the waterways

• amount of biological activity

• location along the catchment (sink, shallow flowing, etc.)

• proximity of sensitive areas or areas of special significance

• variations in the intensity of evapo-concentration

• accessibility to potential sampling locations.

Ideally, there should be a temporal as well as a spatial component to the sampling of temporary waters. Sampling should be distributed evenly along the catchment and through time to get a clear indication of water quality.

C4.2 Sampling release waters Certain considerations need to be taken when sampling temporary water receiving environments after a discharge or release has occurred. Due to the temporary nature of the flow in these water bodies, there is the potential for environmental harm to occur:

• with the resumption of flow, where active ecosystems located downstream of the discharge point are subjected to a pulse exposure to contaminants

• with the cessation of flow, where the concentration of contaminants within remnant pools begins to increase due to the effects of evapo-concentration.

Historical releases during dry phases may have resulted in the accumulation of contaminants in the sediments. With the resumption of flow (either due to rainfall, wastewater release or other cause), these contaminants can be remobilised and may cause environmental harm to ecosystems located downstream. The level of environmental harm (if any) is dependent on the concentration of the contaminants and the level of dilution afforded by the flow. This is discussed in greater detail in section C4.3.1 below.

Sampling of temporary waters at or downstream of the release point should always be accompanied by reference site sampling. Additionally, wherever possible, sampling should take place at multiple points within both the receiving and the reference environments so that suspect or outlier data can be recognised.

C4.3 What to sample There are three main approaches to establishing water quality in temporary waters, namely:

• surface waters

• sediment

• biological assessment.

A brief outline of each of these approaches is presented below. Further guidance to each of these can be found in ANZECC/ARMCANZ (2000).

C4.3.1 Surface waters

The sampling of temporary waters is usually event-based, occurring upon resumption of rainfall during the flood stage of the hydrocycle or release/overflow of waters from an industry. It may take the form of manual grab samples of surface waters, or be performed by remote automatic samplers that trigger sampling at a particular water level or at pre-determined intervals. To ensure adequate monitoring of contaminants, water samples should also be collected as the drying phase of the hydrocycle progresses. Protecting the ecosystem in temporary waters is usually easier during high flow events due to the effects of dilution. However, when flow diminishes, evapo-concentration of contaminants in surface waters can occur, thereby potentially heightening the effects of exposure on components of the ecosystem. It is during the drying phase that many of the ecosystem components are in their reproductive stages. Even in the ‘dry’ state there is an array of live organisms in the sediments, often in desiccation-resistant life stages ready to repopulate the surface waters when the waters return.

It is recommended that the Queensland AusRivAS methodology be adopted when sampling non-permanent waterways (see section 4.1).

It should be noted that the initial flush of water after dry periods will be very likely to give a skewed assessment of

water quality. The first flush that occurs immediately after rainfall may be more damaging (poorer water quality) than the subsequent flow; however, this pulse is typically only short lived and is unlikely to harm encysted or dormant stream fauna.

There are two scenarios that are likely to be encountered where temporary waters exist. If the receiving environment is:

• an ephemeral or temporary waterway without pools, or temporary or permanent water bodies of ecological significance for many kilometres downstream, any live biota still existing within the boundaries of the watercourse will most likely be in a dormant stage below the sediment surface. Consequently, there will be minimal exposure to the first flush of water that enters that watercourse. Hence sampling of remnant water that will harbour and support the emergent biota will be much more ecologically relevant

• an ephemeral or temporary waterway with pools, or temporary or permanent water bodies of ecological significance located downstream, and is likely to be impacted by first flush water flows, permanent or recently established water bodies will most likely harbour active life stages of biota. These will be vulnerable to the effects of a pulse exposure of contaminants associated with first flush waters. In this case, sampling of the first flush water and sampling of the residing water in the permanent/semi-permanent water body (post-exposure) would be warranted.

Due to the variability in flows, it can be useful to incorporate automatic samplers for sampling temporary waters. See section 3.1.2 on using automatic samplers.

C4.3.2 Sediments

Water quality should be measured directly and quantified wherever possible; however, this is seldom possible in arid environments. It is more likely that samples will need to be taken during the dry stage of the hydrocycle. Sampling may have to rely partially or entirely on historical deposition of contaminants in the sediment of dry river beds.

Contaminants in sediment samples can be extracted in the laboratory to estimate their potential bioavailability.

Some of the problems inherent in relying on sediment quality are that:

• the relationship of sediment quality to actual water quality is often uncertain, making extrapolation of data difficult

• only sediment bound contaminants will be measured

• sediment is typically heterogeneous and so care must be taken in ensuring appropriate replication and/or composite sampling.

For further sampling information, refer to AS/NZS 5667.12:1999 Guidance on Sampling Bottom Sediments.

C4.3.3 Biological assessment

Biological assessment can be a valuable way to establish changes in water quality. This usually involves long-term monitoring of assemblages of organisms both at the impacted and reference sites. Biological monitoring can give an early indication of reduction in water quality.

Guidance on selecting the best approach and the associated methodologies of each can be found in Section 8.1 of ANZECC/ARMCANZ (2000). Some of the approaches to biological assessment are:

• direct toxicity assessment

• seed propagule banks

• aquatic plants (algae and macrophytes)

• monitoring of hyporheic and epigean fauna.

Some of these biological approaches may require specialist localised knowledge of taxa, and assistance from the appropriate experts may be difficult to procure.

For further guidance, Smith et al (2004) have produced a valuable review of the benefits and flaws inherent in various sampling methodologies in evaluating the impacts of mining in arid and semi-arid regions of Australia.

Further information on applying the ANZECC/ARMCANZ (2000) guidelines to sampling results for temporary waters have been addressed in Batley et al (2003), A Guide to the Application of ANZECC/ARMCANZ Water Quality Guidelines in the Minerals Industry. This publication also provides some extra advice on sampling strategies for temporary waters.

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Appendix C5 Bulk natural water and sediment collection for direct toxicity assessment (DTA) Direct toxicity assessment (DTA) is a useful tool for characterising the combined toxicity of contaminants in a mixture. The toxicological effects of chemicals in combination can be quite different from the effect of each chemical in isolation. By comparing DTA results with the actual or estimated load of contaminants in a discharge, it is possible to develop a range of predictive conditions and potential effects on biota for specific discharge events.

A limitation of DTAs is that it may be difficult to obtain test species that are ecologically relevant (i.e. present) in the catchment being sampled. In such cases, the relevance of DTA results to field situations may be limited due to different sensitivity between species. Nonetheless, the relative toxicity of the discharge to ‘standard test organisms’ can still be very useful in quantifying the environmental risk posed by the activity.

DTAs can be conducted on bulk water samples or extracts from sediments. Contact the analyst to obtain detailed advice on how to collect samples for DTA before collection.

When collecting bulk receiving environment water samples for the purpose of diluting effluent to be used in DTAs, collect the appropriate volume of water according to the testing laboratory’s advice, which should include:

• bottle type

• field storage requirements

• instruction on any preservatives to be used

• instruction on sample security

• instructions for transport.

If the required information is unobtainable prior to collection, follow the guidance provided in this manual.

It is a requirement that physicochemical parameters (pH, conductivity, temperature, ammonia, dissolved oxygen, turbidity and suspended solids) be measured (in the field wherever possible) at the time of collection in accordance with the instructions in this manual, and then again prior to conducting any dilution of the effluent to be tested for the DTA. There may be a requirement for additional parameters to be tested both in the field and prior to using the collected water for effluent dilutions where the time for transport and use of the receiving environment water in the bioassays exceeds 24 hours. Seek advice on this from the department (who will assess this requirement on a case-by-case basis).

Further information on the use of DTA in water quality assessment can also be found in the review by Smith et al (2004), and in the ANZECC/ARMCANZZ (2000) Australian and New Zealand Guidelines for Fresh and Marine Water Quality.

Appendix C6 Contact details for laboratories Despatch to: Name and organisation street address

Notify: Name telephone and fax numbers

email address

Other contacts: Name telephone and fax numbers email address

Physico-chemical analysis—water, sediment, vegetation and animal samples

Microbiological analysis— water, sediment, vegetation and animal samples

Pathological/toxicological examination—vegetation and animal samples

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Appendix C7 Units and concentrations When taking readings from field instruments and test kits, and in interpreting data from sample analysis relative to guideline thresholds and limits in licences and permits, a variety of equivalent and non-equivalent units may be encountered. Take care with units and be consistent in recording readings and interpreting data. To reduce the risk of errors, using standard units of measurement is preferable to generic terms such as ‘parts per million’.

Concentrations may be written in symbols using a variety of forms, for example milligrams per litre as μg/L or as μg.L-1 both of which are equivalent.

Standard units and their abbreviations

Note particularly that units used to express results by field instruments and test kits are not necessarily consistent with those used in compliance requirements or guideline documents, e.g. phosphate as phosphate (PO4) is not the same as phosphate as phosphorus (P), similarly, nitrate as nitrate (NO3) is not the same as nitrate as nitrogen (N), and ammonia as ammonium (NH4) is not the same as ammonia as nitrogen (N). Use the conversion factors listed below.

Kilogram kg 103 Parts per thousand g/kg mg/g g/L mg/mL μg/μL

Gram g 1 Parts per million ppm mg/kg μg/g mg/L μg/mL ng/μL

Milligram mg 10-3 Parts per billion ppb μg/kg ng/g μg/L ng/mL pg/μL

Microgram μg 10-6 Parts per trillion ppt ng/kg pg/g ng/L pg/mL fg/μL

Nanogram ng 10-9 Parts per quadrillion ppq pg/kg fg/g pg/L fg/mL ag/μL

Picogram pg 10-12

Femtogram fg 10-15

Attogram ag 10-18

Conversion factors for concentrations of N and P compounds

Molar units and their abbreviations

Concentrations may also be expressed in molar units (M). A mole is 6.022 x 1023 particles (of nitrogen, or ammonia, or phosphorus, etc.). A mole of anything contains the same number of items.

A molar concentration is the number of moles in a litre of solution, for example, a 1M solution of ammonia as nitrogen (N) contains 14 grams of nitrogen (N) in a litre of solution (14 gN/L).

Using the standard units table above it follows that a 1 mM solution of ammonia contains 0.014 gN/L, and a 1 µM solution of ammonia contains 0.014 mgN/L.

1 mg/L of nitrate (NO3) as nitrogen (N) = 4.43 mg/L of nitrate as nitrate.

To convert: mg(N)/L to mg(nitrate)/L multiply by 4.43

mg(nitrate)/L to mg(N)/L divide by 4.43

1 mg/L of nitrite (NO2) as nitrogen (N) = 3.29 mg/L of nitrite as nitrite

To convert: mg(N)/L to mg(nitrite)/L multiply by 3.29

mg(nitrite)/L to mg(N)/L divide by 3.29

1 mg/L of ammonia (NH3) as nitrogen (N) = 1.21 mg/L of ammonia as ammonia

To convert: mg(N)/L to mg(ammonia)/L multiply by 1.21

mg(ammonia)/L to mg(N)/L divide by 1.21

1 mg/L of ammonium (NH4) as nitrogen (N) = 1.29 mg/L of ammonium as ammonium

To convert: mg(N)/L to mg(ammonium)/L multiply by 1.29

mg(ammonium)/L to mg(N)/L divide by 1.29

1 mg/L of phosphate (PO4) as phosphorus (P) = 3.066 mg/L of phosphate as phosphate

To convert: mg(P)/L to mg(ammonium)/L multiply by 3.066

mg(ammonium)/L to mg(P)/L divide by 3.066

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Conversion factors for concentrations in molar units

Ammonia – N and Nitrate – N and Nitrite – N

Urea – N

Phosphate – P

Silicon – Si and SiO2

1M ammonia (or nitrate or nitrite) as nitrogen (N) = 14 gN/L,

therefore 1.0 mgN/L ammonia (or nitrate or nitrite) = 71.4 µM

To convert: mgN/L to µM multiply by 71.4

µM to mgN/L divide by 71.4

1M urea as nitrogen (N) = 28 gN/L,

therefore 1.0 mgN/L = 35.7 µM

To convert: mgN/L to µM multiply by 35.7


1M phosphate as phosphorus (P) = 31 gP/L,

therefore 1.0 mgP/L = 32.3 µM

To convert: mgP/L to µM multiply by 32.3

µM to mgP/L divide by 32.3

To convert: SiO2 to Si multiply by 0.4674

1M silica (Si) = 28.1 gSi/L,

therefore 1.0 mgSi/L = 35.6 µM

To convert: mgSi/L to µM multiply by 35.6


Conductivity units and their abbreviations

The unit of measurement for conductivity is siemens (S) per unit of length. A siemen is the reciprocal of the unit of electrical resistance, the ohm (Ω).

Some common EC unit expressions used for reporting conductivity are:

• microsiemens per centimetre (µS/cm or µS.cm-1)

• millisiemens per centimetre (mS/cm or mS.cm-1)

• decisiemens per metre (dS/m or dS.m-1)

• millisiemens per metre (mS/m or mS.m-1).

The SI unit is mS/m. Equivalence relationships among these units include:

• 1 dS/m = 1 mS/cm = 100 mS/m = 1000 µS/cm

• 1 mS/m = 10 µS/cm

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Appendix C8 Sample containers and preservation methods IMPORTANT: Use section 3.3 to interpret these tables correctly.

Table C8.1 Alphabetic list of water quality characteristics covered in this manual

Parameter Quick reference

Acidity and alkalinity

Adsorbable organic halide (AOX)

Aluminium See ‘metals’

Ammonia See ‘nutrients’

Anionic surfactants See ‘surfactants, anionic’

Arsenic See ‘metals’

Barium See ‘metals’

Biochemical oxygen demand (BOD)

Boron

Bromide

Cadmium See ‘metals’

Calcium See ‘metals’

Chemical oxygen demand (cod)

Chloride

Chlorine, free

Chlorine, free residual See ‘chlorine, free’

Chlorine, total

Chlorophylls

Chromium See ‘metals’: separate entries for ‘hexavalent’ and ‘total’ forms

Cobalt See‘metals’

Colour

Conductivity

Copper See ‘metals’

Cyanide

Dissolved oxygen

Electrical conductivity See ‘conductivity’

Enterococci See ‘bacteria’


Escherichia coli and thermotolerant coliforms See ‘bacteria’

Faecal coliforms This term has been superseded by ‘thermotolerant coliforms’

Fluoride

Hardness See ‘metals’—‘calcium’, and follow instructions under that heading

Herbicides See ‘herbicides and pesticides’

Hydrocarbons, petroleum

Iodide

Iron See ‘metals’

Lead See ‘metals’

Lignins and tannins

Magnesium See ‘metals’

Manganese See ‘metals’

Mercury See ‘metals’

Molybdenum See ‘metals’

Nickel See ‘metals’

Nitrate See ‘nutrients’

Nitrite See ‘nutrients’

Nitrogen, oxidised See ‘nutrients’: ‘oxidised nitrogen’ is the sum of nitrate and nitrite

Nitrogen, total See ‘nutrients’

Nitrogen, total kjeldahl See ‘nutrients’

Non-ionic surfactants See ‘surfactants, non-ionic’

Oil and grease

Organotins (e.g. TBT)

Oxygen, dissolved See ‘dissolved oxygen’

Pesticides See ‘herbicides and pesticides’

pH value

Phosphorus, dissolved See ‘nutrients’

Phosphorus, total See ‘nutrients’

Polychlorinated biphenyls (PCB)

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Polynuclear aromatic hydrocarbons (PAH)

Potassium See ‘metals’

Selenium See ‘metals’

Silver See ‘metals’

Sodium See ‘metals’

Solids, dissolved

Solids, suspended

Sulphate

Sulphide

Surfactants, anionic

Surfactants, non-ionic

Suspended solids See ‘solids, suspended’

Thermotolerant coliforms See ‘bacteria’

Total dissolved solids See ‘solids, dissolved’

Trihalomethanes and haloacetic acids

Turbidity

Uranium See ‘metals’

Vanadium See ’metals’

Zinc See ‘metals’

When a parameter is not identified in this manual, contact the analyst for proper collection techniques.

Table C8.2 Water samples—containers and preservation methods Container codes: V, Volume: Typical sample volume, mL (see note [1]). W, Washing: A = Acid, D = Detergent, S = Solvent, W = Water. M, Material: G = Borosilicate Glass, P = Plastic (examples: Polyethylene, PTFE,

[‡] Except where specifically stated otherwise, put sample in container first, THEN add the preservative/s. [§] ‘Refrigerate’ = cool to 1-4°C; ‘Freeze’ = freeze to -20°C. [†] Denotes recommended maximum time from collection in the field to analysis in the laboratory.

Preservation procedure Container code(as above) How much: (a) OR (b)Quality characteristic

(water samples) V W M

Filling method and/or

sample filtering (if any)Preservative

to add [‡] (a) Mass or volume

(b) to pHshown

Storage conditionsRec. max. holding period

Other instructions

cidity and alkalinity 250 D P Fill container completely to exclude air

None — — Refrigerate 24 hours Analyse in the field if practicable (consult analyst about procedure)

Adsorbable organic halide (AOX)

1000 A G Fill container completely to exclude air

Nitric acid

— 1-2 Refrigerate in the dark 3 days Transport sample promptly to laboratory so that analysis can be started as soon as practicable

Bacteria Escherichia coli — — — —

Maximum 24 hours

see note [3]

—

Biochemical oxygen demand (BOD)

1000 W G or P

Fill container completely to exclude air.

None — — Refrigerate in the dark 24 hours Check with the laboratory on suitable arrival time

Boron 100 D P Fill container completely to exclude air

None — — — 1 month —

Bromide 100 D P — None — — Refrigerate in the dark 1 month — Chemical oxygen demand (COD) (use 1 of the 2 methods)

Method 1

100 A G or P

Fill container completely to exclude air

Sulphuric acid

— 1-2 Refrigerate 7 days —

Chemical oxygen demand (COD) (use 1 of the 2 methods)

Method 2

100 A G or P

Fill container completely to exclude air

Sulphuric acid

— 1-2 Freeze 1 month —

Chloride 100 D P — None — — — 1 month —

Chloride, free 100 D P — None — — Keep sample out of direct sunlight

5 min Determine immediately on site

Chloride, total 100 D P — None — — Keep sample out of direct sunlight

5 min Determine immediately on site

Chlorophyll use 1 of the 2 methods)

Method 1

See note [4]

— — See note [4]—keep filter paper (filtered liquid is not required for this test)

None — — Refrigerate in the dark 24 hours Sample must be transported in the dark

Chlorophyll use 1 of the 2 methods)

Method 2

1000

D G or P

— — — — Wrap folded filter paper in aluminium foil and freeze in the dark

1 month —

Colour 500 D P — None — — Refrigerate in the dark 48 hours —

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Conductivity use 1 of the 2 methods)

Method 1

100 W P Fill container completely to exclude air

None — — — 24 hours Determine in situ or on site if possible

Conductivity use 1 of the 2 methods)

Method 2

100 W P Fill container completely to exclude air

None — — Refrigerate 1 month —

Cyanide, Total see note [5]

500 W P — Sodium hydroxide solution(see caution*)

— =>12 Refrigerate in the dark 24 hours *Caution: this procedure can create lethal HCN gas

Dissolved oxygen (DO) use 1 of the 2 methods)

Method 1

300 W G — None — — — — Determine in situ or on site

Dissolved oxygen (DO) use 1 of the 2 methods)

Method 2

300 W G — Winkler solution — — Store in the dark 24 hours —

Fluoride 200 D P — None — — — 1 month PTFE (poly-tetrafluoroethylene) containers are unsuitable

Herbicides and pesticides: Herbicides

1000 S G* Do not pre-rinse container with sample material. Do not completely fill sample container (leave head space of approx. 1-2 cm depth)

Sodium thiosulphate; see note [6]

80 mg per L sample

— Refrigerate in the dark 7 days * Lid of sample container must have insert of aluminium or PTFE (poly-tetrafluoroethylene). Protect sample from light

Herbicides and pesticides: Carbamates

yrethrins ynthetic pyrethroids rganochlorine, rganophosphorus and itrogen-containing esticides



80 mg per L sample

— Refrigerate 7 days * Lid of sample container must have insert of aluminium or PTFE (poly-tetrafluoroethylene)

Hydrocarbons, petroleum 1000 S G Do not pre-rinse container with sample material. Fill bottle completely with sample (no head space)

Sulphuric acid

— 1-2 Refrigerate 1 month —

odine 500 D P — None — — Refrigerate in the dark 1 month — ignins and tannins 250 W G — None — — Refrigerate 7 days —

Metals calcium or magnesium (use 1 of the 2 methods)

Method 1

250 A P Fill container completely to exclude air

Nitric acid

— 1-2 — 1 month —

Metals —calcium or magnesium (use 1 of the 2 methods)

Method 2

250 A P Fill container completely to exclude air

None; see note [7]) — — — 7 days —

Metals —chromium, hexavalent (see note [8])

100 A P — None — — Refrigerate 24 hours —

Metals —mercury

Note: Glass container required, and two preservatives involved.

250 A G For ‘dissolved’ form, filter on site; see note [9]

Nitric acid

and

Potassium dichromate 50 mg/mL

—

5 mL per 250 mL sample; or more. (See right-hand column)

1-2

—

— 1 month If sample is wastes or contaminated waters, more potassium dichromate may be needed: • If sample is initially clear or a pale colour, adding 5 mL of preservative should turn it a distinct yellow. If the colour appears but then fades, more preservative is needed; so add a further 5 mL. Repeat if necessary. • If sample is initially a dark colour, it may be difficult to determine whether enough preservative has been added. Discuss with the analyst before sampling if practicable. Both preservation steps are required

Metals —potassium or sodium use 1 of the 2 methods)

Method 1

100 W P — None — — — 1 month —

Metals —potassium or sodium use 1 of the 2 methods)

Method 2

250 A P Nitric acid — 1-2 — 1 month Permits measurement with other metals

Metals —others: Aluminium, arsenic, barium, cadmium, chromium—total (hexavalent + trivalent), cobalt, copper, iron, lead, manganese, molybdenum, nickel, selenium, silver, uranium see note [8]

250 A P For ‘dissolved’ form, filter on site; see note [9]

Nitric acid

— 1-2 — 1 month —

Nutrients: Ammonia (use 1 of the 2 methods)

Method 1

100 W P Filter on site; see note [9]

None — — Refrigerate 24 hours Store in area free of contamination (ammonia vapour may permeate the walls of containers even if, made of high density polyethylene)

Nutrients: Ammonia (use 1 of the 2 methods)

Method 2


None — — Freeze 1 month Store in area free of contamination (ammonia vapour may permeate the walls of containers even if, made of high density polyethylene)

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Nutrients: Nitrate (use 1 of the 2 methods)

Method 1

100 W P Unfiltered sample None — — Refrigerate 24 hours —

Nutrients: – Nitrate

(use 1 of the 2 methods)

Method 2


None — — Freeze 1 month —

Nutrients: NITRITE (use 1 of the 2 methods)

Method 1

100 W P — None — — — 24 hours Have sample analysed as soon as possible after collection

Nutrients: Nitrite (use 1 of the 2 methods)

Method 2

100 W P — None — — Freeze 48 hours —

Nutrients: Nitrogen, Total Kjeldahl (TKN) or Total (TN)

use 1 of the 2 methods)

Method 1

250 W P — None — — Refrigerate 24 hours —

Nutrients: Nitrogen, Total Kjeldahl (TKN) or Total (TN)


Method 2

250 W P — None — — Freeze 1 month —

Nutrients: —phosphorus (dissolved)

se 1 of the 2 methods)

Method 1


None — — Refrigerate 24 hours —

Nutrients: —phosphorus (dissolved) se 1 of the 2 methods)

Method 2


None — — Freeze 1 month —

Nutrients: —phosphorus (total) use 1 of the 2 methods)

Method 1

250 W P — None — — Refrigerate 24 hours —

Nutrients: —phosphorus (total)


Method 2

250 W P — None — — Freeze 1 month —

Oil and grease 1000 S G Do not pre-rinse container with sample material. Fill bottle completely with sample (no head space)

Sulphuric acid

— 1-2 Refrigerate 1 month —

Organotins (e.g. TBT) 1000 S G — None — — Refrigerate 7 days * Lid of sample container must have insert of aluminium or PTFE (poly-tetrafluoroethylene). Protect sample from light

H 100 D P — None — — Refrigerate 6 hours Determine in situ or on site if practicable

Polychlorinated biphenyls (PCB)



80 mg per L sample

— Refrigerate in the dark 7 days * Lid of sample container must have insert of aluminium or PTFE (poly-tetrafluoroethylene)

Polynuclear aromatic hydrocarbons (PAH)

1000 S G Do not pre-rinse container with sample material. Do not completely fill sample container (leave head space of approx. 1-2 cm depth)

Sodium thiosulphate ; see note [6]

80 mg per L sample

— Refrigerate in the dark 7 days —

Solids, dissolved 1000 D P Fill container completely to exclude air

None — — Refrigerate 24 hours —

Solids, Suspended 1000 D P — None — — Refrigerate 24 hours — Sulphate 200 W P — None — — Refrigerate 7 days — Sulphide 500 D P — Zinc acetate (10%

solution) 2 mL per 500 mL sample

— Refrigerate 7 days —

Surfactants anionic 500 [10] G — Sulphuric acid

— 1-2 Refrigerate 48 hours Glassware must not have been washed with detergent previously

Surfactants, non-ionic 500 [10] G — Formaldehyde 25 mL per L sample

— Refrigerate 1 month Glassware must not have been washed with detergent previously

Trihalomethanes and haloacetic acids (use 1 of the 3 methods)

Method 1

200 S G* Fill container completely to exclude air

Ascorbic acid, or

0.125 g per 200 mL sample

— — 14 days *Glass vial with cap having PTFE-faced septum


Method 2


Sodium thiosulphate; see note [6], or

16 mg per 200 mL sample


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Method 3


Ammonium chloride 0.2g per 200 mL sample


Turbidity 100 D P — None — — — 24 hours If practicable, determine in situ or on site

Table C8.3 Sediment samples – containers and preservation methods Container codes: Typical sample volume, mL (see note 1). A = Acid, D = Detergent, S = Solvent, W = Water. G = Borosilicate Glass, P = Plastic (examples: PTFE, polyethylene, PET, polypropylene or similar) [§] ‘Refrigerate’ = cool to 1-4°C; ‘Freeze’ = freeze to -20°C. [†] Denotes recommended maximum time from collection in the field to analysis in the laboratory.

(SEDIMENT SAMPLES) V W M to add (a) Mass or Volume

(b) to pHshown

period

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Note [1] The volume shown is a typical volume for a single determination. The actual volume needed depends on many factors. If practicable, discuss with the analytical laboratory before sampling.

Note [2] Sterile containers are required as specified in AS/NZS 2031 2001. For samples from chlorinated sources, these must contain sodium thiosulphate before sampling. The concentration must be such as will produce a concentration of at least 100 mg/mL in the sample. Where bottles are prepared by the laboratory, they should be supplied already containing the required amount of sodium thiosulphate. Sodium thiosulphate neutralises the chlorine, thus preventing further bactericidal effects on organisms in the water during transit.

Note [3] AS/NZS 2031 2001 indicates that microbiological examination should commence within six hours of collection; under exceptional circumstances this may be extended to a maximum of 24 hours. The analyst should attach a note to the test report stating the time interval between sample collection and testing.

Note [4] Filter the sample and submit the filter paper itself (along with the particulate matter accumulated on it) to the analyst. Filter the sample in the field using vacuum filtration equipment. Filter the sample through a fine glass fibre filter (Whatman GF/C or equivalent). The vacuum applied across the filter should be no more than 40 cm of mercury, to avoid possible rupture of cells and consequent release of chlorophyll. The volume of sample needed will depend on the concentration of particulate matter present; a typical volume to filter is 1000 mL, but for samples with high loadings, the filter can become clogged before this volume has passed through. You must record the volume that has actually passed through the filter, and give this information to the analyst with the sample. Filters and collected particulates must not be touched with fingers and all sample handling apparatus must be kept free of acids as this causes degradation of chlorophyll. Place filter paper in a container that excludes light (for example, a small tube completely wrapped in aluminium foil) for transport to analyst.

Note [5] When requesting analysis, specify which form you require to be determined.

Note [6] Preservative is needed only if sample is chlorinated. Preservative should be in the bottle prior to filling with sample.

Note [7] Second method (no preservation) can be used only where sample is: [a] of pH = < 8 and low carbonate content; and [b] drawn solely for determination of calcium, magnesium or hardness.

Note [8] When collecting a sample for determination of chromium (VI), it is suggested that you also collect a sample for determination of total chromium and submit both for analysis. This allows the laboratory to perform the simpler test for total chromium first; then, if none is detected, it need not carry out the more complex test for hexavalent (VI) chromium.

Note [9] Filter sample in the field, through 0.45 µm poly ether sulphone filter, preferably using fully enclosed pressure filtering equipment, such as single use syringe (50 mL) with 0.45 µm single use filter. For turbid samples a glass fibre pre-filter should also be used to make filtering easier.

Note [10] Containers to be methanol rinsed glass in accordance with AS 5667.1:1998.

Note [11] You must specify which metals are to be determined in the sample.

Note [12] If analysis is required for total organic carbon (TOC) as well as for pesticides and herbicides, the same sample will suffice for both analyses.

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Appendix C9 Fluvial sediment sampling using P 61 sediment samplers and Helley–Smith bedload samplers

C9.1 Skills/competency and experience Staff skills, training and experience records should be kept up to date. Skills/competency ideal for this method include:

• all members of the sampling party having a current Senior First Aid certificate

• at least one member of the sampling party having had previous training and experience in the use of this method.

During sampling, not all samples retrieved are acceptable. For example, water samples that are contaminated with bedload must not be sent for suspended sediment analysis as they are not representative. Therefore, experimentation may be required to get an idea of the stream condition before taking samples. The whole sampling exercise demands skill, experience and patience and it is highly recommended that a trial run be conducted prior to the wet season for the following reasons:

• ensure that all equipment is operational

• fine tune and improve the data collection exercise

• identify possible problems and seek solutions

• reinforce the tasks to be carried out by each crew member

• provide hands on training for any new crew member.

C9.2 Equipment Equipment specific to this method include:

• Helley-Smith bedload sampler with sample weighting system

• P 61 suspended sediment sampler with current meter

• ADCP in conjunction with a modified van Dorn sampler

• See Box C9.1 for further equipment requirements.

Two different sediment samplers are involved, one for the bedload sediment (Helley-Smith) and the other for the suspended sediment (P 61). The bedload weighing system is another piece of equipment required and is used in conjunction with the bedload sampler for determining the transport rate. This section details the operation of the Helley-Smith bedload sampler, P 61 suspended sediment sampler and the bedload weighing system. The latter is used in conjunction with the Helley-Smith to determine the bedload transport rate.

Sampler location on boats:

P 61 on the starboard side (right) and the bedload sampler on the port side (left).

BOX C9.1 Equipment requirements for fluvial sediment sampling

Description Quantity

Bedload sampling

Bedload sampler

Sampling bags (minimum)

Bags for storage of samples

Bedload weighing system

Bucket for submerged weighing (20 litres)

Wire to attached sampling bag to load cell

Stop watch

Battery 12V 5.7AH

Pencil

Screw driver for attaching sampling bags (if sampler not modified)

Standard (existing) winch

1

5

10*

1

1

1

1

2

4

1

1

Suspended sediment sampling

Suspended sediment sampler

Sampling bottles (minimum)

Plastic bottles for storage of samples

Battery pack (if required)

Spare fuse for battery pack (if required)

Spare nozzle

Modified hanger bar

C2 connector

Modified powered winch

Thermometer

1

24

100*

1

3

2

1

1

1

1

Other Items

Data recording sheets (minimum)

12V truck batteries and battery charger

Generator

Trolley or wheelbarrow (if required)

Crane (if necessary)

Chain for securing crane to bridge

Safety vests

Safety cones

Flashing light

Tomahawk or wire cutters (for cutting four core cable)

Large container for storing samples

12

2

1

1

1

1

3

4

1

1

2

* depending on the number of samples to be collected

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C9.3 Method Sediment sampling involves not only the collection of samples but other related parameters as well. Samples/measurements to be taken are listed as follows:

• point velocity measurement at every location where suspended sediment sample is obtained

• suspended sediment sample

• bedload sample

• timing for each sediment sample (in seconds)

• submerged weight of each bedload sample

• water temperature.

All suspended sediment and bedload samples are to be sent to the Queensland Health Forensic Scientific Services (QHFSS) (or other approved laboratory) for analysis.

C9.3.1 When to measure

Sediments are mainly transported and deposited during flood events. The load is known to be higher during the rising limb than the recession, the rising limb being associated with the removal of stored sediments in the stream or the initial flush of mobile sediment from the land surface, particularly after a severe drought. Stream velocities, and thus sediment transport rates, are also usually higher for a given water level during the rising limb, than during the falling limb.

It is recommended that at least two sets of data be obtained at each sampling site for a flood event, one on the rising limb and the other on the recession limb. Ideally, the measurements should be taken during the period close to the maximum discharge. Figure C9.1 illustrates the two possible starting times, A and B.

Figure C9.1 Possible starting times (A and B) for sampling (for short duration floods)

In most cases, however, the selection of the starting times is very much dependent upon when the hydrographic team arrives and the time required to set up the equipment, particularly in a catchment of quick response. Figure C9.2 shows the period T, where the measurements can be taken.

Figure C9.2 Period (T) where samplings should be conducted

Measurements should cease when it takes a long time to collect the bedload (e.g. greater than about 10 minutes). For floods of longer duration (e.g. two or more days), advantage should be taken to collect more than two sets of data. In this case the selection of the starting times for sampling can be better decided. Figure C9.3 presents the possible starting times, A to D, for four sets of sediment sampling.

Figure C9.3 Possible starting times for sampling (for long duration floods)

C9.3.2 Selecting verticals for sampling

The number of verticals required in a cross-section for sediment sampling is dependent upon the width, discharge distribution, accuracy sought, as well as the concentration variation across the river at the time of sampling. The location of each vertical is normally chosen based on:

• equally spaced verticals—with this method, the channel width at the water surface is divided into sections of equal width corresponding to the number of verticals required. This is a relatively easy method to use as the discharge need not be known prior

• strips of equal discharge—in this method, verticals are arranged according to the distribution of water discharge across the section. Each sampling vertical represents approximately equal portions of discharge. It is, however, a more involved process as the discharge must be computed prior to the sampling work

• consideration of the transverse distribution of sediment concentration—another method of selecting the verticals is to consider the transverse distribution of sediment concentration. In zones of large concentration variation,

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verticals are placed closer together whereas a lesser number of verticals is used on the floodplain.

Recommendation: Location of verticals should be established before sampling for a possible range of flow depths to avoid additional workload in the field. Consult with the Project Manager.

C9.3.2.1 Flow confined within main channel

Figure C9.4 shows an example of a selection of five equally spaced verticals (or six equal segments) across the stream. Where operationally possible, the distance between each vertical should not be more than 50 metres). This will, however, be stream/site-specific.

Figure C9.4 Location of sampling points (flow confined within main channel)

NOTE: for ADCP measurements, the distance from the bottom and top differs due to back scatter noise.

C9.3.2.2 Over bank flow

In the event of significant over bank flows, more verticals should be placed in the main channel and less over the floodplain. Figure C9.5 presents an example where the main channel is divided into four equal portions and one vertical is placed in the middle of the floodplain.

Figure C9.5 Location of sampling points (significant over bank flow)

NOTE: for ADCP measurements, the distance from the bottom and top differs due to back scatter noise.

However, the selection of the verticals at some sites may differ from the above to meet specific program objectives and advice should be sought from the Project Manager. To minimise the time required for completing a set of sediment measurements (and also possible confusion), the sampling should be carried out as a separate exercise to the stream gauging. The latter requires a larger number of verticals.

C9.3.3 Sampling methods for suspended sediments

There are two methods commonly used for sampling of suspended sediment, namely depth and point integration:

• Depth integration—in depth integration, the water–sediment mixture is taken continuously while the sampler is moving at a constant transit rate throughout a vertical. However, this method is limited to a depth of 4.5 m only for round trip sampling or 9 m for single trip. Furthermore, the transit rate of lowering or raising should be kept constant and should not exceed four-tenths of the mean velocity of a vertical. This condition must be adhered to for reliable results.

• Point integration—point integrated sampling involves an accumulation of water sediment mixture that is representative of the mean concentration at any selected point in a stream, over a short period of time. There is at least one selected point in a vertical.

Recommendation: In view of the operational difficulty and the depth constraint for the depth integration method, the more robust point integration method is recommended.

The number of sampling points in a vertical should vary according to the depth of the stream and the size of sediment in suspension. Commonly used methods are the one, two, three and five point methods. Their relative depths (measured from the water surface) are shown in Figures C9.4 and C9.5. More points selected would imply a longer duration of sampling (which may be counter productive) and result in higher costs for laboratory analyses. Hence the accuracy desired has to be balanced against cost and time.

Recommendation: Locations of the sampling points are recommended as follows:

Flow confined within the main channel

The middle vertical has five points at relative depths of 0.0, 0.2, 0.6, 0.8 and 1.0 The rest of the verticals are taken at relative

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depths of 0.2 and 0.8 (Figure C9.4).

Over bank flow

The five point sample is preferably taken in the deepest part of the main channel. For the rest of the verticals, samplings are taken at relative depths of 0.2 and 0.8 (Figure C9.5).

C9.4 Equipment use

C9.4.1 Acoustic Doppler current profiler

The acoustic Doppler current profiler (ADCP) is a standard instrument used in the collection of stream flow data. During this process, backscatter information from suspended sediment particles is also collected. This backscatter data can be used in the calculation of sediment loads (as water quality samples are collected in conjunction with each ADCP run). However, several points need to be considered before collecting point sediment samples with a P 61 sampler when using the ADCP:

• Blanking distance (blank after transmit)—the same transducer is used to receive the acoustic energy after transmitting a pulse. A short time (or a short sound travel distance) must pass before receiving is possible. This delay is called the blanking distance, and it allows the ADCP to ring down and become acoustically quiet before receiving the return signal. For the 600 kHz and 1200 kHz Rio Grande ADCPs, this distance is 25 cm.

• Near zone distance—there is also a shallow layer of water near the bottom for which the data is not used to compute discharge. When the ADCP sends out an acoustic pulse, a small amount of energy is transmitted in side lobes rather than in the direction of the ADCP beam. Side lobe reflection from the bottom can interfere with the water echoes. This gives erroneous velocities for the water near the bottom. WinRiver II does not use data in the region that may be affected. The ADCP has beams oriented at 20 degrees from the vertical, and the thickness of the side lobe layer is 6 per cent of the distance from the transducers to the bottom (see Figure C9.6).

• Edge estimates—as the name suggests, the flow for the edges is estimated using a power fit method of the data collected near the edge. In most cases for the collection of velocity data for load calculation this would not be an issue.

Figure C9.6 Side lobes Source: Adapted from WinRiver diagram

Due to the methods outlined in Section C5.2, there can be issues due to the effects mentioned above; therefore, some considerations to these effects are required, i.e. modification to the location of the samples (shown in Figure C9.7) to allow for velocities to be extracted from the ADCP data. The operator should be able to quickly calculate the depths required from the output from WinRiver in the field, as the depth at which the ADCP is located (mounted on the boat) and the depth of the water are the influencing factors. The changes required are minimal and any

Bottom surface

94%

6%

Side lobe Main lobe

20 degree beam angle

ADCP

major errors should occur due to the changes in the depth of sample; however, this would require verification before implementation. A detailed operational procedure on ADPC/Sediview applications is to be developed for approval as a standard method.

Figure C9.7 Unmeasured regions in the water column

C9.4.2 Helley-Smith bedload sampler

Various types of bedload sampling equipment have been developed and used throughout the world. The Helley-Smith, a pressure difference type, is most commonly used overseas. This sampler is used for bedload sampling (see Figure C9.8). It has an intake nozzle which measures 76 mm by 76 mm. The unit weighs about 48 kg (105 lb), is constructed of a welded stainless steel assembly and has a stabiliser tail assembly. Bedload is caught in a polyester mono filament bag with a mesh opening of 0.2 mm which will not absorb water.

The sampler is best suited for sampling sediments in the range of 0.5 mm to 16 mm, which is appropriate for a large number of alluvial streams in Queensland. Larger sediments will need to be sampled with a larger sized nozzle (152 mm). The sampler is calibrated in a laboratory to determine its efficiency coefficient, defined as the ratio of the trapped sediment to that actually moved as bedload per unit of time. For the range of applications over medium to coarse sand and gravel bed (0.5 mm to 16 mm), the efficiency is close to 100 per cent for flow velocity of less than 3 m/second.

Figure C9.8 Helley-Smith bedload sampler (labelled A) and P 61 suspended sediment sampler (labelled B)

Sampler assembly

1. Remove the bedload sampler from the storage box.

2. Suspend the sampler in the water and adjust the suspension bracket so that the sampler (tail heavy) is at an

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angle of approximately 25 degrees from the horizontal. To achieve this, release the locking screws and move the sliding collar as an adjustment. Lock the screws and ensure that the suspension bracket is vertical.

3. To reduce the amount of time for set up on site, this step should have already been performed during the trial run prior to the actual measurement.

4. Place the sample bag, with seam uppermost, to the rear of the nozzle.

5. Place an elastic band on the sample bag in the location groove around the nozzle.

6. Slide the metal band over the elastic band, ensuring that the locating pins on the sides are in the secure position for a snug fit.

7. Secure the bag by pulling the clip into a locking position. The metal band and clip arrangement is a modification made to the original equipment for improving handling procedure in the field (original equipment has four clamps which can only be released by a screwdriver—a slow and frustrating job on a boat with a good chance of losing the screws holding the clamps).

8. Connect the hook to the eye at the rear of the sample bag.

9. Connect the C1 connector to the sliding collar.

Sampling

1. Locate the sampling vertical position determined previously.

2. Lower the sampler carefully to the stream bed. When the tail makes contact, slowly lower the nozzle until it is sitting on the bed and start the stop watch; then record the sampler depth. Care should be taken not to dig the nozzle into the stream bed.

3. Collect the sample over a period long enough to give a decent sample (which will depend on the local velocity and bedload particle size). Raise the sampler immediately when the time expires.

4. Retrieve the sample bag. This task is easier to perform if the sampler is hauled into the boat.

2. Ensure that the sample bag is not filled by more than 40 per cent, else discard the sample and repeat the exercise with a reduced duration of sampling.

3. Weigh the wet sample, fully submerged in water and record its weight in grams (g).

4. For samples that require laboratory particle size analysis (PSA), transfer the contents into another container. Ensure correct labelling and record the sample number.

5. Discard the material from the sample and wash out the sample bag.

6. Record other details as required in the data recording sheet.

7. Assemble the sample bag and repeat sampling.

8. It is recommended that two or three readings be taken at each location. If uncertain or results differ markedly, an additional sample is justified.

Dismantling the sampler

1. Remove the sample bag from the sampler for storage.

2. Ensure the lower clamp screws and clamps are secured to prevent loss or damage in the storage box. For recently modified samplers, ensure that the steel band is properly stored.

3. Return the sampler to the storage box for transportation.

For further procedures on sampling with the Helley-Smith bedload sampler refer to the Hydrological Services Operating instructions for bedload sampler model BLS-30 and model BLS-48

C9.4.3 Bedload weighting system

This system is used to measure the submerged weight of the bedload sample. If the sediment sampling is carried out from a boat, it is highly desirable that the weighing be done on the boat itself, rather than saving the samples for weighing later while on land. When weighing is carried out immediately, the hydrographer is aware of the quality of the data and can make an informed decision as to whether another sample should be collected to improve reliability of the results.

Sampler operation

1. Remove the weighing system from the Peli-case container.

2. Assemble the weighing system (refer to schematic in Figure C9.9):

a. Connect the twisted pair wires between the back panel of the digital readout and the battery (e.g. 12V 5.7 AH Sonnenschein).

b. Plug the cable from the load cell into the rear panel of the digital readout.

c. Attach the two hooks to either end of the load cell.

d. Hang one end of the hook to a bar (or a rigid frame).

e. Switch on the digital readout and wait for about 10 seconds for unit to stabilise.

3. Ensure that the readout is set to register in g (not lbs), else press the ‘weight selection’ key.

4. Hang another empty sampling bag (for bedload) on the hook and totally immerse the bag into a weighing bucket part full of water (e.g. 20 L).

5. Zero the digital readout and remove the empty bag.

6. You may need to provide a length of wire so that the sampling bag can be adjusted to allow for full immersion. Recently supplied bags are fitted with hanger lips for direct attachment to the load cell hook.

7. Weigh the collected sample bag and its bedload fully immersed in the bucket of water. Place the bag gently onto the load cell, taking care not to snag the hook instantaneously. Record the weight in g.

8. If the boat is rocking or high wind conditions prevail, press the ‘Autohold’ (second blue button at the bottom of the readout) for 2 seconds. The ‘Held’ bar will flash on the indicator. The display will then search for a steady weight to lock onto. To turn off this mode, press the ‘Autohold’ for another 2 seconds.

9. An overflow occurs for weight in excess of 7000 g. This should not occur under normal operating conditions where the sediments consist of sand and gravel.

WARNING: Do not overstress the load cell by the amount indicated in the manual (20 kg).

10. Remove the sampling bag and sediments from the weighing assembly, discard the contents and thoroughly wash out the sampling bag.

11. The digital indicator weighing assembly does not need to be switched off if constantly in use, else repeat steps 4 and 5. Carry a spare battery for the weighing assembly.

12. Re-attach the emptied and washed sampling bag to the bedload sampler and repeat the exercise.

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Figure C9.9 Schematic of bedload weighing system

In the past, all bedload samples were analysed in the laboratory for their dry weights. However, a weighing technique, based on the work by Carey (1984), is available whereby the submerged weight can be converted to the dry weight from the knowledge of the specific gravity of the bedload material. The relationship is given by:

Wd = [SG/(SG - 1)]Ws

where :

Wd = dry weight

SG = specific gravity

Ws = submerged weight

For quartz material, the SG is equal to 2.65, and hence: Wd = 1.606 Ws

A major advantage of this technique is that the weighing can be conducted in the field, thereby eliminating the time consuming exercise and costs involved in saving all the samples for laboratory analyses. Only samples that require particle sizing are kept and this represents only a very small quantity. This technique allows more samples to be collected and weighed at no additional costs, and hence improves the reliability of the data.

If the sediment sampling is carried out from a boat, it is highly desirable that the weighing be done on the boat itself, rather than saving the samples for weighing later on land. When weighing is carried out immediately, the hydrographer is aware of the quality of the data and can make an informed decision as to whether another sample should be collected to improve the reliability of the results.

This system was specifically designed for submerged weighing with the desired accuracy.

C9.4.4 P 61 sampler and current meter

The purpose of a suspended sediment sampler is to obtain a representative ‘discrete’ sample that is representative of the water–sediment mixture in a stream in the vicinity of the sampler. As such, the device is carefully constructed to satisfy several design criteria and it is also calibrated.

Integrating samplers collect a sample over a period of time to average out concentration fluctuations. The P 61 point integrating sampler for suspended sediment is the most common sampler employed in several countries and is used in this method (Figure C9.8). Developed by the US Geological Survey, the P 61 weighs about 48 kg, has a streamlined body and is cast in bronze with tail fins for orientation with the flow.

A solenoid activated valve opens and closes the nozzle during sampling. The water–sediment mixture enters the sampler into the container through the nozzle as air is exhausted through the air-exhaust opening. As such, the sampler should not be overfilled as the desired flow rates will not be attained and the sample will become contaminated by the flow circulating in through the nozzle and out through the exhaust opening.

Sampler assembly

1. Remove the P 61 from the transport container.

2. Connect the hanger bar to the P 61.

3. Assemble the current meter above the P 61 and lock it into position; then measure and record the distances between the current meter centreline and 1, the bottom of the P 61 (e.g. 0.42 m) and 2, and the P 61 intake nozzle centreline (e.g. 0.32 m).

4. Attach the C2 connector to the top end of the hanger bar.

5. There will be two cables dangling from the C2 connector. Plug one of the cables to the current meter. The other cable plugs into the head of the P 61. A locking lever ensures that the plug is secured.

6. On the winch end there will be four cables. Plug two cables to the readout unit of the current meter, while the remaining cables are connected to the battery pack.

7. Artificially rotate the current meter propeller and to confirm that a readout is registered, else interchange the cables. The cables should, however, be marked as ‘Meter’ or ‘Battery’ for easy identification.

8. Test the operation of the solenoid valve by holding the switch to the ‘charge’ position on the battery pack, until the meter registers a reading of about 48 V. Press the switch to the ‘sample’ position. A click on the P 61 indicates that the valve is operational.

9. operate the winch to raise the P 61 (and current meter) into a working position

10. Open the head and insert the appropriate P 61 sampling bottle (cap removed) with the aluminium adaptor into the body of the P 61. Close the head slowly but firmly to ensure that it snaps into position. It’s most convenient to have 13 pre-numbered P 61 sampling bottles which are capped after sampling, and then decanted into laboratory sample bottles back on the bank after each gauging.

11. The sampler is now ready. Take note of the horizontal distance between the sampler and the centre line of the boat.

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Sampling (point integration)

1. Locate the sampling vertical.

2. Lower the sampler to the streambed to determine the depth of flow.

3. Raise the current meter to the appropriate position for velocity measurement.

4. Record the point velocity to enable determination of sample duration.

5. Raise (or lower) the P 61 by an appropriate distance to the previous position occupied by the current meter.

6. Press the switch on the battery pack to the ‘sample’ position for the desired sampling duration.

7. Release the switch on the battery box and hoist the sampler into the boat.

8. Open the head gently, ensuring that the head is always inclined upwards by at least 30 degrees to prevent spillage of the sample.

9. Remove the sample bottle with care.

NOTE: The 500 mL bottle must not be greater than 2/3 full (or 333 ml). If overfilled, discard the sample and repeat the suspended sediment sample collection with a shorter time period. The recommended volume is between 200 and 300 mL. Both levels should be marked on all sampling bottles prior to the event.

NOTE: Sampling time is dependent on stream velocity, so experimentations may be necessary at the start, or devise a local relationship between sampling time and current meter revs (e.g. 3000/C31 Prop 1 Revs).

10. Label the analysis bottle with the water quality field analysis number. Decant the sample into the analysis bottle for the laboratory or, if necessary, another container (e.g. 1 L bottle). This container can be decanted later to the analysis bottle on return to shore. This step isn’t necessary during sampling if 13 pre-numbered P 61 sampling bottles are used.

11. Replace the sample bottle in the P 61 and close the head.

12. Move to the next sampling position in the vertical and repeat sampling steps 2 to 12 until all points in a vertical are completed.

Possible problems at step 8

The solenoid valve may be seized due to deposits formed from prolonged storage. Connect the power supply directly to the sampler, apply power, and strike the side of the sampler head with a rubber or wooden mallet. Never strike the nozzle.

Another reason for failure could be due to a blown fuse in the battery pack. Open the top of the battery cover and the fuse is located on the circuit board just beneath the cover (see Figure C9.10).

Check to see the fuse is operational. If not, change the fuse and replace the cover.

These additional ‘repair type’ jobs are not desirable during the flood event, thus reinforcing the point that the equipment should be well maintained and a trial run carried out prior to the wet season.

Figure C9.10 Battery pack showing fuse location

13. Move to the next vertical, ensuring that the bedload sampling for the vertical has also been completed.

14. Repeat the process from steps 2 to 14 until all verticals for the cross-section have been sampled.

15. Obtain the temperature of the water (2 m from the bank) if no normal suit of samples are being obtained during the sample run.

16. Back at the bank, decant any samples remaining in P 61 sampling bottles into labelled laboratory sample bottles.

Special care for P 61 samplers

The following care should be taken with the P 61 sampler:

• Handle the nozzle with care; if bent or burred, this will contribute to sampling error.

• Never submerge a sampler without a sample bottle in the body.

• Try not to overfill the sample bottle; otherwise water can flow back through the air exhaust opening into the head.

• Disassemble the head and clean (and dry) all parts after use to prevent the solenoid valve from seizure during storage.

C9.5 Equipment maintenance Sediment sampling presents a more complex exercise compared to the normal velocity measurements as:

• heavier equipment is involved

• the existing winch and cable arrangement has to be modified to accommodate the 4-core cable

• more tasks need to be performed by the hydrographic team.

To ensure a successful program, all equipment needs to be operational and it is vital to maintain it in a good state of repair. A bench test of all equipment is a prerequisite, including assembling the equipment on a boat, crane or vehicle. Other checks are also required such as:

• operation of the bedload sampler

• operation of the suspended sediment sampler

• sampling bags and bottles are in good condition

• accuracy and operation of the bedload weighing system

• modified winches are operational

• connector and hanger bar are in working condition

• battery pack for the suspended sediment sampler is operational

• modifications made to the boat, crane, etc. are in working condition.

C9.6 Sample handling

C9.6.1 Bedload samples

One sample is to be kept for each data set (i.e. one for every field sheet). This sample, from the deepest part of the channel, is sent to the laboratory for sieve analysis. To assist in the identification of the samples, each sample bag is labelled with the field water quality analysis number using a permanent marker. Details of the sample are completed on the field sheet.

C9.6.2 Suspended sediment samples

The top and bottom suspended sediment samples from the deepest point are analysed for particle size analysis and sediment concentration. The remaining three suspended samples from the deepest point and the other verticals are to be sent for sediment concentration type analysis. The quantity of suspended sediment samples requiring labelling is therefore greater than the bedload samples.

C9.7 Quality assurance Complete and accurate completion of field sheets is necessary.

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Control all possibility of sample contamination. Refer to Part B of this manual.

C9.8 References AS/NZS 5667.11:1998, Water quality—Sampling: Guidance on sampling of groundwaters, Standards Australia.

Carey, W.P. (1984) A field guide for weighted bed load samples, Water Resources Bulletin, American Water Resources Association, Vol 20, No. 2.

Teledyne Technologies website <www.rdinstruments.com> site last accessed 7 April 2009.

Wong, W.T., Alexander, D.G. and Eades G., Proceedings from the sediment sampling workshop, October 1992.

Appendix C10 Sampling procedures for suspended solids and nutrients—application of water sampling technique NOTE: The procedure in this appendix includes the triple-rinsing of sample containment bottles with site water prior

to taking the sample. This is NOT appropriate when taking samples using specially-prepared bottles such as acid-washed or solvent-washed bottles. Nor is it appropriate if the sample bottles are supplied with the sample preservative already in the bottle (rinsing would lose the preservative). If a bottle for sample containment is supplied pre-cleaned by the analysing laboratory, no useful purpose is served by rinsing it on site. Although it is best practice to use specially-cleaned bottles supplied by the analysing laboratory for sample containment, new bottles, not specifically prepared may be used for sampling nutrients and suspended solids, and in this circumstance, triple-rinsing is appropriate.

C10.1 Background for sampling procedures Sampling involves the collection of a water sample directly into a suitable sampling container (direct collection) or collection into a temporary sampling container (intermediate container) with subsequent transfer into recommended bottles for storage. Appropriate containers and preservation methods are necessary to avoid risks of contamination of the sample and/or losses of analytes of interest during storage and transit prior to analysis. Details of appropriate sample containers and preservation methods are given in Appendix C8.

When sub-sampling from an intermediate container or when multiple sub-samples are drawn from a single sample, it is important to ensure the sample is homogenous prior to sub-sampling. The use of a sample splitting device (such as a churn splitter or similar) is recommended to achieve this.

There are a number of ways that samples can be collected and the choice of method will be determined by the analyte/s of interest, sampling objectives, and site specific conditions. Equipment used for intermediate and direct sample collection can include the following:

1. an extendable rod and bottle holder that allows for the sample bottle to be manually filled from the stream at a distance away from a stream bank (see section 3.1.1)

2. United States Geological Survey type isometric depth sampling equipment and/or a rope and weighted discrete sampling vessel (e.g. van Dorn sampler) for sampling from bridges, boats or overpasses

3. an automated pumping sampler (see section 3.1.2 Automatic Samplers) allows samples to be pumped from a point in stream to an intermediate sample containers (some refrigerated) for collection. Samplers can be triggered remotely or at pre-specified times or river heights

4. rising stage samplers (RSS) allow samples to be collected from a range of water depths with sample bottles filled as the river level rises during a flow event. Samples can only be collected on the rise of a flow event and are retrieved after the river level has receded (see section 3.1.1).

The choice of sampling equipment is largely dependent on the purpose for which the data is collected. Refer to section 2.1 for guidance on when and where to sample when applying these methods.

When collecting samples using any of these methods it is necessary to prevent contamination of samples (see section 3.5.1.).

Guidance on methods of sample collection provided below is of a general nature. As there are many different types of isometric samplers and van-Dorn like vessels, automatic, and rising-stage samplers available, methods may need to be modified slightly. In such cases, manufacturer instructions should be used for guidance. For all methods, once samples are collected, samples should be preserved and stored in bottles and as appropriate for each analyte of interest (see Appendix C8) and as guided by program objectives.

C10.2 Manual sampling procedures Where it has been deemed appropriate and safe to collect a discrete unit of water, a sample may be collected manually using an extendable pole sampler or an isokinetic, discrete depth sampler if appropriate.

C10.2.1 Sampling using extendable pole sampler

1. Remove the lid from the 1 L sample bottle and attach the bottle to the end of a sampling rod.

2. Extend the sampling rod into the main flow of the stream. Submerge the bottle to a depth of at least 0.3 m, keeping the mouth end pointing down.

3. Whilst submerged, rotate the sampling rod 180 degrees to bring the mouth of the bottle facing up, and allow the bottle to fill with water; and retrieve bottle.

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4. * Replace the lid and shake the bottle ensuring the inside of the bottle and the lid come into contact with the liquid. Discard the rinse liquid downstream of where you are sampling. Be sure to keep hands away from the mouth of the bottle and the underside of the lid.

5. *Repeat steps 3 to 4 so that the sample bottle and its lid are rinsed twice with stream water then proceed to step 6.

6. *Repeat step 1 to 3 to fill the bottle. Replace the lid and tighten. If sample requires freezing, ensure you leave 10–20 per cent space free.

*These additional steps (bottle rinsing) only appropriate if specifically-cleaned bottles are not being used to collect samples.

C10.2.2 Manual sampling using isokinetic and discrete depth samplers

For methods to operate sampling for specific isokinetic samplers and van-Dorn like vessels, see manufacturer’s instructions.

1. Lower the sampling device into the main flow of the stream to a depth of at least 0.3 m.

2. Whilst submerged, trigger the opening/closing device as per manufacturer instructions and allow the bottle to fill with water.

3. Remove the device from the water and agitate to resuspend solid material.

4. Open device and pour into storage bottle in a smooth and constant movement keeping materials in suspension.

5. Replace the lid on storage bottle and shake the bottle ensuring the inside of the bottle and the lid come into contact with the liquid.

6. Discard the liquid downstream of where you are sampling.

7. Repeat steps 4 to 6 so that sample device and storage bottle and its lid are rinsed twice with stream water, discard remaining water downstream.

8. Lower the sampling device into the main flow of the stream. Submerge the device to a depth of at least 0.3 m.

9. Whilst submerged, trigger the opening/closing device as per manufacturer instructions and allow the bottle to fill with water.

10. Remove the device from the water and agitate to resuspend solid material.

11. Open device and pour into storage bottle in a smooth and constant movement keeping materials in suspension.

12. Fill storage bottle, replace the lid and tighten. If sample requires freezing, ensure you leave 10–20 per cent space free

Figure C10.1 Van Dorn sampler

C10.3 Automatic sampling procedures Automatic samplers should be loaded with sample containers that are prepared using the necessary methods to avoid risks of contamination of the sample and/or losses of analytes of interest during storage and transit prior to analysis. Details of appropriate sample containers and preparation and preservation techniques are given in Appendix C8.

For guidance on sample bottle installation and sampler operation please refer to manufacturer instructions.

C10.3.1 Processing samples from automatic pumping samplers

1. Place lids on auto-sampler (intermediate) bottles whilst in the carousel.

2. Remove auto-sampler bottles individually in a sequential order from the auto-sampler carousel (Figure C15).

3. For each auto-sampler (intermediate) bottle follow steps 4 to 7.

4. Shake sample vigorously to resuspend settled material.

5. Remove lid from auto-sampler bottle and rinse storage sample bottle by carefully decanting a small amount of sample into the storage bottle. Shake and ensure the inside of the storage bottle and its lid comes into contact with the liquid then discard rinse water.

6. Repeat this process so that the bottle and its lid are rinsed twice with sample water.

7. Replace the lid on auto-sampler bottle and again shake vigorously to resuspended settled sediment then quickly decant sample into sample bottle for storage.

Figure C10.2 Removing samples from the auto-sampler

C10.3.2 Processing samples from rising stage samplers

For guidance on sample bottle installation and sampler operation please refer to manufacturer instructions. Note that the sample inflow assembly of rising stage samples require rinsing prior to use.

Once river levels have receded enough to allow safe access to the rising stage sampler proceed with sample collection.

1. Remove the in-flow assembly of one bottle from the rising stage sampler, starting from the top and secure sample bottle with a clean, dry lid; if the sample bottle used is an intermediate container please follow steps 4 to 7 in C10.3.1.

2. Label the sample bottle clearly indicating what its position in the rising stage sampler. Start numbering from the bottom up as this is the order in which the bottles filled in (i.e. number in the reverse order to the order of removal).

C10.2 Sampling procedures for filtered nutrients For any of the sampling methods above, the following steps should be followed when collecting filtered nutrient samples.

1. Refer to instructions for sample collection above. Once sample has been collected:

2. From the sample bottle take up 10 mL in a 60 mL syringe then fully extend the plunger and shake liquid to rinse the syringe ensuring all the inside surface comes into contact with the sample. Expel the water from the syringe downstream from the area where you are working. Repeat to rinse the syringe twice.

3. Completely fill the syringe with sample water from sample bottle and attach a filter (and pre-filter if required) to the end of the syringe as shown below (Figure 7a,b).

4. Discard the first 2 mL of the sample water pushed through the filters as a filter rinse (where possible) and then continue pushing another 10 mL of the water sample into the sample bottle.

5. Replace the lid on storage bottle and shake to rinse bottle and its lid. Repeat so that the storage bottle and its lid are rinsed with filtered sample water twice.

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(a) (b)

(c)

Figure C10.3 Attaching the pre-filter (a) and filter (b) to the syringe and filtering sample into bottle ‘E’ (c)

1. Remove filter and fill the syringe completely to 60 mL with sample water from sampling bottle, re-attach filter and use this to fill appropriate storage bottle (Figure 7c). If this becomes difficult, change the filter and/or pre-filter making sure you rinse at least 2 mL of sample water through them before filtering the water into the storage bottle again to the required analysis volume. Make sure to leave at least 10–20 per cent free space for water expansion. Secure the lid and store sample according to appropriate preservation techniques for later analysis.

PART D Sampling bio-indicators of water quality and environmental health

4.1 Macro-invertebrate sampling and assessment This methodology is based on the Queensland AusRivAS (Australian River Assessment System).

4.1.1 Introduction to AusRivAS The structure of plant and animal communities of rivers can give us a far more accurate picture of the condition or health of our waterways than measuring water quality parameters alone. Of these biological communities, macroinvertebrates (i.e. animals without backbones and large enough to be seen with the naked eye, e.g. prawns, shrimps, crayfish, snails, mussels and insects such as dragonflies, damselflies and mayflies) are most widely used because they are abundant and diverse, and are sensitive to changes in water quality, flow regime and habitat conditions. Impacts on these animals are relatively long lasting and can be detected for some time after the impact occurs. Impacts on a waterway can be varied: chemical spills, riparian vegetation removal, sand and gravel extraction, or stock access. All of these things can upset the balance, e.g. a chemical spill might kill a large proportion of the macroinvertebrate community, which is a major component of the diet of some fish, and consequently the fish population affected.

The AusRivAS (Australian River Assessment Scheme) model protocol (Simpson et al. 1997) adapts the River Invertebrate Prediction and Classification Scheme (RIVPACS) methods applied by Wright et al. (1984), Moss et al. (1987), Marchant et al. (1994) and aspects of protocols developed by Chessman (1995). This approach to assessing river health was adopted by the National River Health Program (NRHP—comprising the Monitoring River Health Initiative (MRHI) and First National Assessment of River Health (FNARH)). It allows rapid sampling methods to be used for the development of predictive models for macroinvertebrate communities within each state/territory, using a ‘reference’ site database. Comparisons may then be made between predicted and observed taxonomic compositions of macroinvertebrate communities in different habitats at a site in order to indicate the presence and magnitude of an impact on the site’s ecological health. This approach can assess biological responses to changes in water quality and/or habitat condition in rivers and can be integrated with the existing network of physico-chemical water quality monitoring sites.

The actual protocol used (e.g. sampling design, number of sites, subsampling, replication, frequency of sampling, etc.) should be based on the objectives of the monitoring or assessment program. The AusRivAS protocol outlined below is meant for broad scale monitoring (e.g. catchment or regional basis). For smaller scale and specific issues, other methods using control and replicate sites (such as BACIP and multivariate equivalents) may be more appropriate (see Underwood 1993; ANZECC 2000).

The development of a standardised tool such as AusRivAS for broad scale bioassessment is dependent on three factors:

• use of the same biota

• use of the same approach to sampling and sample processing

• use of the same analytical methods for model development and use.

The data can also be analysed in different ways and used for other purposes such as impact assessments, condition and trend reporting, biodiversity and biogeographic studies.

It should be noted that this protocol is for use only in freshwater reaches of rivers and not for use in estuaries or tidal reaches of lowland rivers. Although the general approach may be valid, substantial additional work must be performed prior to its adoption in estuarine and marine conditions.

4.1.2 Sampling program Important note:

The sampling program described below was followed in developing the AusRivAS predictive models for Queensland rivers. Anyone intending to utilise the models for assessing riverine sites must follow this procedure. Data can then be input into the models run by EHP. Those not intending to use the EHP models but still interested in developing and implementing a broad scale biological monitoring and assessment program for their streams and rivers can also follow this procedure. The advantage of doing so is that a standard method is followed which would permit direct temporal and spatial comparisons of data, and allow sites to be compared with existing MRHI and FNARH reference and test sites.

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4.1.3 Site selection Reference and test sites in Queensland were initially selected for the MRHI program using protocols outlined in the River Bioassessment Manual (Davies 1994). Reference sites were those ‘least disturbed’ sites sampled for the production of a database used in the construction of predictive bioassessment models (e.g. RIVPACS of Wright et al. 1984). Test sites were those identified to be of importance in assessing the condition of a river known or perceived to be experiencing an impact from water quality or habitat degradation. These protocols were adapted by QNR&W into a list of criteria to which each site was subjected. Table 4.1 lists the 10 selection criteria currently used to determine whether sites are in reference condition.

The input of additional information from outside sources may potentially result in modifications to these assessments. This method will provide a better characterisation of the sites based on numerical categorisation bringing the assessments into line with other programs undertaken by QNR&W. Each criterion relates to an aspect of human activity that impacts on freshwater ecosystems, where impact is defined as a ‘change from natural condition’. Each criterion is given a score according to the following categories:

1. very major impact

2. major impact

3. moderate impact

4. minor impact

5. indiscernible impact.

Table 4.1 Selection criteria used to determine eligibility for reference site status No. Reference condition selection criteria

1 Influence of intensive agriculture upstream.*

Intensive agriculture is that which involves irrigation, widespread soil disturbance, use of agrochemicals and pine plantations. Dry-land grazing does not fall into this category.

2 Influence of major extractive industry (current or historical) upstream.*

This includes mines, quarries and sand/gravel extraction.

3 Influence of major urban area upstream.

This will be relative to population size, river size and distance between the site and the impact.

4 Influence of significant point-source waste water discharge upstream.*

Exceptions can be made for small discharges into large rivers.

5 Influence of dam or major weir.*

Sites within the ponded area of impoundments also fail. Sites failing this criterion automatically fail the overall assessment.

6 Influence of alteration to seasonal flow regime.

This may be due to abstraction or regulation further upstream than the coverage by criterion 5. Includes either an increase or decrease in seasonal flow.

7 Influence of alteration to riparian zone.

Riparian vegetation should be intact and dominated by native species.

8 Influence of erosion and damage by stock on riparian zone and banks.

Stock damage to the stream bed may be included in this category.

9 Influence of major geomorphological change on stream channel.

Geomorphological change includes bank slumping, shallowing, braiding and unnatural aggradation or degradation.

10 Influence of alteration to instream conditions and habitats.

This may be due to excessive algal and macrophyte growth, by sedimentation and siltation, by reduction in habitat diversity by drowning or drying out of habitats (e.g. riffles) or by direct access.

* Note: the level of impact at a site will generally decrease with the distance from the source of impact

Sites are assessed using the total score for the 10 criteria. Currently, those sites that have a total greater than 44 are deemed to be reference sites. Sites that are given a score of 1, 2 or 3 for criterion 5 (no dam or major weir upstream) cannot be reference sites.

4.1.4 Sampling frequency Any program that attempts to standardise sampling protocols across Australia must take into account the occurrence of rivers encompassing a wide range of predictability and seasonality in river hydrology, life histories of biota and river habitat types. Whether a strong seasonality occurs or not, a minimum number of samples must be taken over time to allow collection of adequate macroinvertebrate taxa information for reference site classification.

The overall aim of the sampling protocol is to ensure that the broadest range of biota are captured at a site by sampling a number of habitats and on a number of occasions. To ensure standardisation and compatibility of data sets, the following protocol was followed for all sites, whether reference or test sites.

The sampling protocol for Queensland rivers and streams requires a minimum of two sample sets in one year. These are sampled on a ‘seasonal’ basis from October to December (early wet—when flow has been established

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for at least four weeks) and May to July (late wet—recessional baseflows when flow had declined to a sampleable level, without significant flood peaks). The early wet samples are identified as ‘spring’ samples; the late wet as ‘autumn’ samples.

For the model development, each site was sampled twice in one year and the data from the two sample sets were used separately to develop seasonal models, and combined to develop an annual model.

4.1.5 Habitats sampled Each reach of a stream may have several habitats. If a habitat accounts for more than 10 per cent of the stream reach then it should be considered for sampling. The predominant habitat types are identified at each site and appropriate ones sampled separately. In Queensland, only two habitats are sampled, an edge sample and a bed sample. The first choice of bed habitat is a riffle; failing this a rocky bed is sampled and, finally, a sandy bed. Ensure that the type of bed sampled is recorded on the field sheet.

Separate sampling of distinct habitat types is prescribed because each habitat has a potentially distinct fauna. The performance of the predictive models will therefore not be confounded by differences in habitat availability between sites and times. AusRivAS models have been developed for edge and pool habitats in Queensland for autumn and spring seasons, as well as a combined season annual model. Riffle models may be developed in the future.

In Queensland, the habitats most likely to be encountered are as follows.

4.1.5.1 Riffle

This is a reach of relatively steep, shallow (less than 0.3 m), fast flowing (more than 0.2 m/s) and broken water over stony beds (see Figure 4.1).

Figure 4.1 Riffle habitats

4.1.5.2 Run

A run is a reach of relatively deep and fast flowing, unbroken water over a sandy, stony or rocky bed (see Figure 4.2). These are features of streams during a flood event, below dams where riffles have been ‘drowned’ or in steep gradient streams flowing through gorges. Under normal flow conditions, it is best to avoid sampling this type of reach. However, pools and riffles may become runs during flood events (e.g. in April) and it may be necessary to sample the area. If possible, delay sampling until flow recedes.

Figure 4.2 Run habitat

4.1.5.3 Pool bed

Pool bed habitats are zones of relatively deep, stationary or very slow flowing water over silty, sandy, stony or rocky beds. This habitat occurs in the main channel and should not be confused with backwaters, which occur as indentations of the bank. Waterholes are generally pools with silty/sandy beds while pools with rocky/gravel beds are often found in steep areas. The velocity will indicate whether it is a pool or run. The classification factor is the bed type, i.e. sandy/silty beds and rocky/gravel beds (see Figure 4.3).

Figure 4.3 Sandy bed and rocky bed habitats

4.1.5.4 Edge/backwater

Edges (or banks and underbank areas) are along the bank where there is little or no current and extend to approximately 0.5 m from the bank. There may be some terrestrial vegetation (e.g. paragrass, sedges) or tree roots, or the area may be bare (e.g. waterholes in drier areas). A backwater is a zone where the bank indents and a pool of water forms away from the main channel (e.g. ox-bow, off-cut channel). The backwater may have a circular or back flow, and a silty bed with accumulated plant litter (leaves, twigs. etc.) (See Figure 4.4).

Figure 4.4 Edge habitats

4.1.5.5 Macrophytes

Macrophyte habitats are areas where emergent, submergent and floating macrophytes or aquatic plants are present and can occur in slow to fast flowing areas. Macrophytes that you are likely to encounter include milfoil (Myriophyllum spp.), hornwort (Ceratophyllum demersum), waterfern (Azolla spp.), salvinia (Salvinia molesta), duckweed, water thyme (Hydrilla verticillata), water primrose (Ludwigia spp.), water hyacinth (Eichhornia crassipes), waterlilies (Nymphaea spp.), pondweeds (Potamogeton spp.) and ribbonweed (Vallisneria spp.). These areas are designated on the field data sheet as ‘macrophytes’ (see Figure 4.5). Although reference and some test site sampling has been done, it is not envisaged that a model will be developed for this habitat. Therefore, this habitat is no longer sampled as part of the AusRivAS approach, although it may still be sampled for other purposes, e.g. nutrient enrichment studies and flow assessments.

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Figure 4.5 Macrophyte habitats

4.1.6 Preparing for a field trip Before embarking on a field trip, some preparation is needed. Appendix H1 and Figures 4.6 and D4.7 outline a list of the equipment needed for macroinvertebrate sampling.

Figure 4.6 Monitoring and sampling gear

Figure 4.7 Monitoring and bug picking gear

4.1.7 Field sheets Prior to water or macroinvertebrate sampling, information must be recorded on field sheets about the site. This includes information about the whole reach (100 m section of the river), the habitats sampled, and the surrounding terrestrial environment. There are four types of field sheets used by EHP to record information about a site:

• Queensland Site Information Sheet (Appendix H2)

• Water Quality Sampling Field Sheet (Appendix H3)

• Habitat Assessment Field Sheet (Appendix H5)

• Task Sheet (Appendix H6).

Note: All EHP field sheets contain common essential information:

• Site number: All sites are allocated a site number, based on the catchment and subcatchment and whether it is a hydrographic gauging station or a ‘miscellaneous’ site. These numbers contain seven alphanumeric characters: the first three numbers indicate which catchment the sites are in (e.g. 136); the fourth digit, the subcatchment; and the final three, the site. For example, a site with the site code 136017B shows that this site is in the Burnett River catchment (136), within the 0 subcatchment, and the site number within this subcatchment is 17B. Sites with a letter as the final character indicates that these sites have or have had a flow gauging station nearby

• Site name: describes where the site is on the river, e.g. Burnett River at Gayndah Flume.

4.1.8 Water quality sampling All water quality measurements and water samples are required to be collected upstream of the biological sampling area and of the water sample collector. They should be taken from a representative section of the stream, slightly flowing if possible, at a depth of 10–20 cm. Care must be taken to avoid sampling too close to the edge and too close to aquatic plants. Field alkalinity measurements are taken using a titration kit (Figure 4.6). Appropriately calibrated meters are needed to measure temperature, conductivity, dissolved oxygen, pH and turbidity (Figure 4.7). The results are recorded on the Water Quality Sampling Field Sheet (Appendix H3).

The water quality samples are collected in prepared bottles, details completed on the sample bottles, samples preserved correctly, external paperwork completed and samples cross-referenced on the Water Quality Sampling Field Sheet. EHP routinely collects two water samples: one for major ions analysis and one for nutrient analysis (nitrogen and phosphorus). Refer to Part B of this manual for appropriate sampling methodology.

4.1.9 Biological sampling

4.1.9.1 General considerations

Sampling should not be conducted when streams are in flood unless the impact of flood is being investigated. If, during the scheduled sampling period, sites are consistently in flood, sampling should resume 4–6 weeks after floods have subsided. The study site is a 100 m length of stream (50 m upstream and 50 m downstream of the point of entry).

All macroinvertebrate samples should be collected with a standard 250 µm mesh dip net. Recommended dimensions are a triangular 250 mm x 250 mm x 250 mm opening and 50–75 cm depth, and with a 1–1.5 m aluminium handle. The net should be checked for damage prior to a sampling trip and washed thoroughly after sampling each habitat to remove animals left from previous sampling. Sample a total distance of 10 m, covering a variety of velocities, if possible, and different examples of the habitat. (Nets are available from the Australian Centre for Tropical Freshwater Research, James Cook University.)

4.1.9.2 Sampling the habitats

Sampling the riffle

While holding the net downstream with its mouth facing the sampling area, disturb the substratum by digging the foot well into the stones and turning them over. Turn and rub stones by hand to dislodge organisms. Continue this process working upstream over a total distance of 10 m, covering both the fastest and slowest flowing sections of the riffle. Do not include material from macrophytes and/or wood debris located in the riffle. It may be necessary to collect the sample from more than one riffle if the first riffle is less than 10 m in length.

Sampling the pool/bed:

Disturb the substratum by kicking with your feet. If the stream is flowing, hold the net downstream with the mouth facing the disturbed area. If there is no discharge you will have to use a short sweeping action with the net while stirring up the bed. The suspended benthic animals are captured as the net sweeps through the cloud of suspended matter.

Note: Silty/sandy beds—preferably select an area with plant litter (not macrophytes) rather than an area of clean sand.

Note: Rocky/gravel beds—if the rocks are too large to kick over without damaging your foot, wash about 10 rocks of a range of sizes, scrubbing gently with the hands or a light brush into the net. Leave the rocks out of the water to allow cryptic specimens to emerge. These can then be hand picked. Be careful, as leaving the rocks in the sun for too long will dry out and kill the animals. Again, avoid areas where macrophytes are present.

Sampling the edge

Locate an edge area with little or no current or aquatic vegetation (stands of paragrass are acceptable as edge

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habitat). An alcove or backwater with abundant benthic leaf litter is preferable. Suitable areas include fine organic/silt deposits and/or trailing vegetation and are often indicated by the presence of surface-dwelling insects. In waterholes you may have no choice but to sample the bare edges, perhaps with some tree roots. Using short upward sweeping movements at right angles to the bank, sample a total bank length of 10 m. Stir up the bottom while doing so, ensuring that benthic animals are suspended and then caught when sweeping through the cloud of suspended material. There may be aquatic plants (macrophytes) along the banks and in backwaters. Avoid sampling these areas.

Sampling the macrophytes:

Locate an area with dense aquatic vegetation (if present). Vigorously sweep the net within the aquatic vegetation over a length of 10 m. Aim to sample the upper, middle and lower portions of the plants. A combination of short lateral sweeps with vertical lifts will aid in dislodging and catching suspended organisms. (This habitat was not modelled for AusRivAS.)

4.1.9.3 Picking the sample

Nationally, two methods are used for collecting organisms: field picking and laboratory picking. The choice of which method to adopt will be influenced by considerations of the objective of the study, precision required, time, cost and balance of effort in the field versus laboratory. Either method may be used, although it is preferable to maintain the same technique for all sites. For Queensland, the field picking option has been adopted as the preferred standard method.

Field picking is considered to be more subjective. However, with sufficient training, care and objectivity, it does provide a cost-effective alternative to laboratory picking.

Field picking:

Treat samples from each habitat separately unless it is part of the project design to have composite samples. It is recommended that the sample is initially separated into two fractions (the small organic and substrate material and large rocks and leaf litter) using a 1 cm panning sieve (cheap aluminium ones are available from local camping stores). Sort through these fractions, a small amount at a time, retaining the residue for QA/QC requirements (see below). Work progressively through the sample, replacing picked material with remaining parts of the sample as picking progresses.

Half fill a vial with 70 per cent alcohol (methylated spirits). Ensure the container you use is large enough. If the animals you collect take up more than 30 per cent of the volume, use a larger container. Use alcohol stable vials to avoid vial cracking and sample loss.

Pick for a minimum of 30 minutes, using tweezers and pipettes, and record the total abundance using a hand held counter.

Collect only 10 of any one type (family and, in some cases, order) of animal. If you are not sure of the identity, then collect all of the uncertain ones. At least 30 midge larvae (Chironomidae) should be collected to ensure adequate representation of the sub-families.

At the start of your field pick, the common and abundant taxa should be picked for about the first five minutes. After that, the major picking effort should be directed at finding the less common, inconspicuous taxa. After 10 minutes no more common taxa should be picked unless it is suspected that a particular common form contains more than one family, or it was a common taxon overlooked initially.

If you get 200 animals (about 10 of each type plus at least 30 Chironomidae) then stop at the end of this 30 minute period. If, at the end of the 30 minutes, you have not collected 200 animals then you should collect for a further 10 minutes. If any new taxa are found in those 10 minutes, extend the picking time by another 10 minutes. Follow this procedure until either no new taxa are found, 200 animals have been collected or 60 minutes have been spent on picking. Note the picking time on the field sheet.

Particular care should be taken to search for the groups that can be commonly missed when live sorting (cryptic taxa):

Corbiculidae (juveniles) Oligochaeta (including broken fragments)

Chironomidae (larvae and pupae) Elmidae (larvae)

Empididae (larvae) Hydrophilidae (larvae and adults)

Hydroptilidae (larvae) Simuliidae (larvae)

Ceratopogonidae (larvae)

If it is a really poor sample (i.e. urban tributary or sandy stream) with very few animals in total, then stop at 60

minutes. Make it clear on the field sheet that it was a poor quality site or sample and why that is so. A poor sample may also result from a bad collection, e.g. a sample taken during high flows over areas which were dry a few days before.

If it is raining or cold, or conditions of poor light exist due to cloud cover or approaching twilight, the sample must be taken back to the vehicle, motel, camp, etc. for sorting undercover and with improved light conditions.

Ensure a completed label is placed in the vial noting the project name, site number and name, sampling date, habitat sampled, sample collector and picker and any relevant notes (see Figure 4.8).

Remove some of the diluted alcohol in the vial using a mesh-covered syringe and replace with fresh 70 per cent alcohol. Fill to top and ensure the lid is tightly screwed on.

Figure 4.8 Label for picked sample

Laboratory picking:

The entire sample is preserved in the field using 70 per cent methylated spirits, a completed label included and the sample adequately stored for transport to the laboratory. Large plastic screw-top jars or heavy-duty plastic bags stored in a polydrum are suitable containers.

4.1.9.4 Field picking quality assurance/quality control (QA/QC)

The residues of 10 per cent of all samples taken in the field are retained for analysis. The entire residue is preserved in the field using 70 per cent methylated spirits, a completed label included and the sample adequately stored for transport to the laboratory. Half of these samples are put aside for external analysis; the other half are subsampled and 10 per cent of each sample are analysed by the unit’s staff. The data is analysed, compared to the sample picked in the field, and reports written.

4.1.9.5 Handy tips for field picking

Use waterproof paper for field sheets.

Only use pencil to fill in field sheets and sample labels; ink and felt tip pens smudge and run.

Some macroinvertebrate groups are fairly cryptic, i.e. difficult to detect, particularly in the first 15–30 minutes after collection. To counteract this phenomenon and ensure that you pick a representative sample from the habitat, it is suggested that all the habitats be sampled before picking begins.

Remember to label each sample as you put it into the tray or bucket. The first sample will then have had sufficient time to rest and the animals will have become more active and easier to see (although not necessarily easier to pick). With labels:

• place them inside the vial

• preferably, DO NOT use gummed (stick on) labels

• if you have to use gummed labels, DO NOT remove the backing paper (the animals stick to the label and are irretrievable).

Buckets are always useful—to carry rinsing water, to put your rock collection in, and to split your sample if it is too big. Ensure that you take at least two per person, preferably more. A couple of buckets with lids are also useful if samples need to be transported before picking.

For safety reasons, at least two people should conduct the field sampling. Field time can be best utilised with one person conducting the macroinvertebrate and water quality sampling and the other filling out the field sheets. Once the samples are collected, these can be live picked in the field by both persons to save time.

4.1.10 Laboratory macroinvertebrate sample processing It is recommended that a registration system be set up for all samples collected. A standard form is recommended

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for this purpose. These sheets should be filled out for all samples within 24 hours of return from field trips. Cross-checking should be performed so that samples recorded on field sheets are present, labels are accurate and legible and bottles filled with preservative. Labels should have the site name, location code, habitat, sampling method, collector’s name, picker’s name and date on them (Figure 4.8).

All registration sheets should be filed appropriately and samples stored in labelled lidded containers (to minimise evaporation) in an approved fire-proof storage area. Each box should contain a logical group of samples to facilitate sample identification and handling.

Software for sample registration and archiving should enable integration with the sample database (see below).

4.1.10.1 Sample identification and enumeration

4.1.10.1.1 Field-picked samples

Ensure adequate ventilation in the workplace. Rinse the sample with gently running water through a 250 µm sieve. Flush the sieve contents into a large petri dish with water from a squeeze bottle. Always use water when working with the sample. When finished, replace the water with preservative.

Place the petri dish under a stereomicroscope which is correctly adjusted for your vision and work posture (refer to manufacturer’s instructions). Use a vial of suitable size to take the collection of specimens in the petri dish with label inserted. The label should have the following information: collection number, location code, site name, collection date, habitat, sample identifier and the identification date (Figure 4.8). Use pencil or alcohol-proof ink to fill in details. Half fill the vial with 70 per cent ethanol.

A dedicated tally sheet (Appendix H7) should be developed for recording the identities and numbers of all taxa in a sample. The sheet should allow listing of the taxonomic key used for identification for each family, the person making the identification, the site, date and sample code.

Organisms are identified to family level with the exception of lower Phyla (Porifera, Nematoda, Nemertea, etc.) Oligochaetes (freshwater worms), Acarina (mites), and microcrustacea (Ostracoda, Copepoda, Cladocera) for which family level identification is optional (it may improve resolution but is time consuming). Chironomids should be identified to sub-family level. Appendix H8 lists the keys used by EHP.

Select specimens and follow the appropriate taxonomic keys to family level. If uncertain about the identity obtain a second opinion from a colleague/local specialist. If a new family is suspected or other significant issues arise in taxonomic identification, contact the relevant national taxonomic specialist.

Identify each specimen, place in the vial and mark the tally sheet. When all specimens have been counted, record the total tally for each taxon. Place the vial, filled with preservative, in an evaporation-proof container (e.g. a large screw-top glass jar) in a suitable storage location.

Transfer the collected data to an electronic spreadsheet or database. At the completion of the sample series, the database should be cross-checked against the data sheets to ensure that there are no transcription errors.

All sorted collections should be archived, preferably lodged with a regional or state/territory museum, so that any future taxonomic revisions or more detailed identifications can be conducted if required. The relevant museum staff should be consulted well in advance of submission of specimens.

4.1.10.1.2 Fully preserved samples

Tip the preserved sample into a series of 10 mm and 250 mm sieves and thoroughly wash the sample. If there are large coarse fractions (sticks, leaves, etc.) wash these over the sieves and place them into a sorting tray. Examine these coarse fractions preferably using a magnifying glass, for approximately 10 minutes, ensuring that any macroinvertebrates attached to the coarse fractions are collected. Note: keep an eye out for stick and leaf-cased Trichoptera.

Evenly distribute the remaining smaller fractions from the sieves into a subsampler. The subsampler used and recommended by EHP is a modified Marchant subsampler (Marchant, 1989) (see Figure 4.9). This subsampler contains 100 circular cells, each 3.5 cm in diameter x 3.5 cm deep. Fill the subsampler until the water level reaches the top of the cells, secure the lid, and rotate vigorously in both directions until the sample is distributed throughout the cells. Using a vacuum pump, subsample 10 per cent (10 cells) of the whole sample, ensuring every 1 per cent (1 cell) of the subsample is stored in separate containers.

Sort and identify the subsamples in the procedure described above (see live picking), noting how many new taxa there are in every 1 per cent subsample sorted. If this procedure is used for internal QA/QC checks on field picking, only 10–15 organisms of each family are identified and recorded. However, if the subsampling is required for quantitative sampling, sort and identify all taxa from the subsamples.

Figure 4.9 Modified Marchant subsampler

The long-term aim in the development of this protocol is to further identify samples to genus and/or species level, in order to improve the predictive power of the resultant models. Thus a high emphasis should be placed on the development of a systematic and well designed data entry and sample archiving system. For certain objectives such as impact assessments, it is advisable to identify the animals to the lowest possible taxonomic level, preferably species, although, at present, AusRivAS models are available only for family level assessment. Identifications made at species/genus level can be converted into family level data and run through the models, but the converse is not possible without re-identification.

4.1.10.2 Laboratory identifications QA/QC

Internal QA/QC checks on laboratory identifications are performed on staff, by staff, on a regular basis. At each round of QA/QC, a person is assigned to analyse a sample identified by another. Samples identified during the previous fortnight are selected at random and re-identified. The resultant taxa lists are compared and discrepancies in identification checked by other staff in the unit. Any errors are discussed with the original identifier (both misidentifications and errors of enumeration) and a report prepared which is read and signed by all members that underwent the QA/QC check. Under NRHP guidelines, error rates greater than 10 per cent in identification and counting are not acceptable.

4.1.11 Database entry and software support A dedicated database should be established to allow for entry, storage, checking and distribution of data and for manipulation of data for subsequent analysis. No specific software package is recommended, although consideration should be given to the ultimate size of the database, the need for ongoing addition of data as the protocol develops into a usable tool, and compatibility with existing systems and software. The database must allow data files to be translated into ASCII format. Data can be entered by a standard spreadsheet (e.g. Excel) and exported to the database (e.g. ACCESS).

Anyone wishing to utilise the AusRivAS model must comply with data formatting indicated in the AusRivAS Manual (visit <www.ausrivas.canberra.edu.au> or Simpson et al. 1997). The appropriate predictor variables should also be considered. Appendix H9 lists the predictor variables used for the Queensland models.

4.1.11.1 Biological data

An appropriate data entry spreadsheet has been designed as recommended by Simpson et al. (1997). Enter all sample data into a unique file and all sets must match exactly. When data is entered it must be cross-checked against data entry sheets. Use the national taxonomic codes for all families (Appendix H10). Use of this coding will allow compatibility between data sets at the agency, state/territory and national levels and will greatly expedite comparisons and future taxonomic efforts.

4.1.11.2 Habitat data

Design a data entry spreadsheet. Perform all calculations to the raw data to conform with the data requirements for later analysis. Enter this processed data for a site into a unique file. Cross-check the data against field sheets. Use

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the same site code and the relevant date and sample coding to allow integration of habitat data with the biological samples for later statistical analysis.

4.2 Blue-green algae (cyanobacteria) sampling and assessment

4.2.1 Introduction to blue-green algae Planktonic cyanobacteria, or blue-green algae, are a common, naturally occurring and integral component of many aquatic ecosystems. As such they often cause no obvious ecological problems. However, a small group of genera produce toxins (cyanotoxins) that have caused sporadic cases of animal poisoning, have been implicated in human hepatoenteritis, and have led to human fatalities.

Cyanobacteria pose a potential health risk through the consumption of water supplies contaminated by cyanotoxins, and through direct exposure to cyanotoxins during water-based recreational activities (i.e. skiing, bathing, wading, boating, etc.). Cyanotoxins have been isolated, identified, and characterised from a number of cyanobacterial taxa, and the ones reported from Australian freshwater to date are presented in Table 4.2.

To reduce the risks of adverse health impacts caused by cyanobacteria, it is essential to routinely monitor water supplies for their presence on a quantitative basis and, where necessary, test for the presence and concentration of their toxins.

Table 4.2 Potentially toxic cyanobacteria reported from Australian freshwaters

Toxic species Cyanotoxin Target organ

Anabaena circinalis saxitoxins nervous system Aphanizomenon ovalisporum cylindrospermopsin liver Cylindrospermopsis raciborskii cylindrospermopsin liver Microcystis aeruginosa microcystins liver Nodularia spumigena nodularins liver Nostoc cf. linkea unidentified liver

4.2.2 Guidelines for assessing blue-green algae The Australian Drinking Water Guidelines (NH&MRC/ARMCANZ 1996) currently provide guidance for one of the four major classes of cyanotoxins found in Australian freshwaters. The guidelines recommend that the concentration of total microcystins in drinking water not exceed 1.3 µg L-1 (expressed as microcystin-LR toxicity equivalents). Due to the lack of adequate data, no guideline values have been set for the other three classes of cyanotoxins. Given the presence of cyanotoxins other than microcystins in Queensland freshwaters, EHP recommends a conservative approach be adopted wherein a value of 1.0 µg L-1 is used for cylindrospermopsin and Paralytic Shellfish Poisons (PSPs) as an interim guidance level until sufficient data has been collected to allow for the development of individual guidelines for these two cyanotoxins.

The risk of adverse health effects through recreational contact in water containing cyanobacteria varies with the concentration of cyanobacteria and the nature of the activity. The risk is derived from contact with both the known cyanotoxins produced by the species listed in the World Health Organization (WHO) Guidelines (Table 4.3), as well as from a group of yet unidentified, non-specific compounds produced by cyanobacterial taxa which have been recorded as producing symptoms ranging from mild skin irritations to nausea, and pneumonia-like symptoms.

Table 4.3 WHO guidelines for safe practice in managing bathing waters which may produce or contain cyanobacterial cells (after Chorus & Bartram 1999)

Hazard status Guidance level or situation Health risks Recommended action High Cyanobacterial scum formation in

contact recreation areas or > 100 000 cells total cyanobacteria mL-

1 or > 50 µg L-1 chlorophyll-a with dominance of cyanobacteria or > 12.5 mm3 L-1 cyanobacterial biomass

Short-term adverse health outcomes such as skin irritations or gastrointestinal illness following contact or accidental ingestion Severe acute poisoning is possible in worst ingestion cases

Immediate action to prevent contact with scums Signs to indicate HIGH alert level—warning of danger for swimming and other water contact activities

Moderate 20 000–100 000 cells total cyanobacteria mL-1 or 10–50 µg L-1 chlorophyll-a with dominance of cyanobacteria or 2.5–12.5 mm3 L-1 cyanobacterial biomass

Short-term adverse health outcomes, e.g. skin irritations, gastrointestinal illness, probably at low frequency

Signs to indicate MODERATE alert level—increased health risk for swimming and other water contact activities

Low < 20 000 cells total cyanobacteria mL-1 or < 10 µg L-1 chlorophyll-a with dominance of cyanobacteria or < 2.5 mm3 L-1 cyanobacterial biomass

Short-term adverse health outcomes unlikely

Cyanobacteria either absent or present at low levels—continue monitoring

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The provisional guidelines for cyanobacteria in bathing water as adopted by EHP are presented in Table 4.4. These guidelines are based on the WHO guidelines for safe practice in managing bathing waters which may produce or contain cyanobacterial cells (Chorus & Bartram 1999). Rather than providing a single threshold value, these guidelines are framed as a series of three guidelines, which reflect incremental severity and probability of adverse effects. These guidelines can be assessed using cell concentrations or cell biovolume concentrations. The later measure is preferred, as it takes into account the huge size range of cyanobacterial cells that occur in natural populations and recognises that the contribution of cyanobacterial cells to the hazard is proportional to their cell volume.

Cyanotoxins have also been demonstrated to pose serious adverse effects on mammals, birds and fish and as such are being increasingly recognised as a potent stress and health hazard factor in aquatic ecosystems. Exposure of aquatic organisms may occur both orally by uptake of toxin-contaminating cells as food, or through the surface tissues of organisms submerged in water containing dissolved cyanotoxins. Despite a growing body of evidence on the ecological effects of these compounds, there are currently no accepted guidelines for cyanobacteria and their toxins relating to the protection of aquatic ecosystems.

4.2.2.1 Storage classification

A national approach to cyanobacterial monitoring taking into account the varying needs and objectives of government agencies, councils, community groups, and members of the public was developed with the creation of a Draft National Protocol for the Monitoring of Cyanobacteria and their Toxins in Surface Waters (Jones et al 2002).

Table 4.4 Outline of monitoring classes for national cyanobacterial sampling protocol (after Jones et al 2002). These are the provisional guidelines adopted by EHP.

It recognised a number of monitoring classes which decrease in effort and cost, and therefore precision and certainty of monitoring and analysis outcome, from that termed ‘class A1’ monitoring, to that termed ‘class C’ monitoring.

The scheme is three tiered, with each tier providing a different level of sampling and analytical precision, and overall certainty of monitoring outcome. Storage operators should determine the appropriate sampling regime based on this monitoring class classification system. A summary of the monitoring classes is given in Table 4.4.

4.2.3 Monitoring sites Buoyant cyanobacteria tend to accumulate near or at the shoreline at the down wind or down stream end of reservoirs or river reaches. Therefore, for high priority public health surveillance monitoring, ‘depth integrated’, open water sampling is preferred. The selection of sampling sites will depend on a number of factors including prevailing winds, the position of stream inflows and the proximity to potential nutrient input sites. Open water sampling provides, in general, a better representation of the true or average cyanobacterial population of the water body. Open water or mid-stream sampling is normally carried out by operations staff working from a boat. For drinking water supplies, sampling the appropriate depth next to, or from the water off-take tower, is desirable.

In general, one open water site in the vicinity of each recreational area, and one sampling site at the water supply off-take tower, should give adequate coverage for both the water supply and recreational use health issues posed

Monitoring class Recommended use category General method description Surety of results A1 Public health surveillance of drinking water

supplies. Condition and trend monitoring—high priority water bodies

Open water sampling from boat Detailed visual surveillance (for scums) High counting precision (< 30% counting error)

High to very high

A2 Public health surveillance of recreational water bodies. Condition and trend monitoring—moderate priority water bodies

Shoreline or bank sampling Detailed visual surveillance (for scums) High counting precision (< 30% counting error)

High

B1 Condition and trend monitoring—moderate priority water bodies

Open water sampling from boat Low to moderate counting precision (> 30% counting error)

Moderate to high

B2 Condition and trend monitoring—low priority water bodies. Community group monitoring

Shoreline or bank sampling Low to moderate counting precision (> 30% counting error)

Moderate

C Public health surveillance for scum formation in bathing waters. Community group monitoring of cyanobacterial growth

Visual surveillance only High (for scum surveillance only) Low

by cyanobacteria. It is desirable that both sites are marked with a buoy or similar device to ensure that sampling occurs in exactly the same area, no matter who samples.

4.2.4 Equipment required for algae sampling The following equipment is required for the sampling of algae:

• integrated hose-pipe sampler—5 m length of 2.5 cm diameter plastic tubing with a weighted collar at one end (see Figure 4.10)

• a cord attached to the hose and boat

• a rubber stopper to fit one end of the tubing

• a bucket

• Lugol’s iodine preservative solution

• 200 mL amber Polyethylene terephthalate (PET) plastic bottle and lid.

Figure D4.10 Procedure for use of the integrated hose-pipe sampler

4.2.5 Sample collection In order to obtain a representative sample for species identification and cell count over the surface depth range, each water sample should be collected at the clearly marked sampling point using the 5 m long, 2.5 cm diameter integrated hose-pipe sampler (Figure 4.10). Sampling should be carried out in the middle of the day, preferably about 1 pm.

The procedure for collecting the sample with a hose-pipe sampler is as follows:

• Attach a cord from the boat to one end of the hose-pipe to prevent accidental loss of the equipment.

• Rinse the sampler by rapidly dropping the weighted end of the hose-pipe vertically into the water to a depth of approximately 5 m while holding the hose-pipe at the top end, and then returning the hose-pipe to the boat without inserting the rubber stopper.

• Again, while holding the hose-pipe sampler at the top end, rapidly drop the weighted end of the hose-pipe vertically into the water to a depth of approximately 5 m. This time, insert the rubber stopper into the top end of the hose-pipe after it reaches its full depth of immersion.

• Pull the bottom end of the hose-pipe to the surface using the cord, so that the tube is in a U-shape (see Figure 4.10).

• Lower the weighted end of the hose-pipe into a clean bucket and remove the rubber stopper. Ensure that the entire contents of the hose-pipe are emptied into the bucket. Mix the contents of the bucket and then transfer

insert rubber stopper

weighted collar

retrieve sampler in a U‐shape

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part of the contents into a 200 mL amber PET plastic bottle, leaving a 25 mm gap at the top of the bottle. The remainder of the contents of the bucket may be discarded.

Algal scums should not be included in the water sample for routine identification and enumeration; however, if they are present, a note should be taken indicating their nature and extent. This is particularly important in bathing or water recreation areas. Additional scum samples can be collected and submitted for qualitative analysis if extensive.

NOTE: Blue-green algae can cause skin irritation. If sampling from an area that has a high level of blue-green algae, minimise your contact with the water during sampling by wearing appropriate dress, in particular gloves. Normal hygiene precautions such as washing off any splashes and washing hands before eating or drinking should be observed at all times. When not in use, the hose-pipe sampler and bucket should be kept clean and stored in a dark shed or cupboard.

4.2.5.1 Sample storage and preservation

To ensure the sample remains in a condition suitable for identification and enumeration, a sufficient volume of Lugol’s iodine preservative solution should be added to the sample to render the sample a colour resembling weak tea (i.e. 1.0 mL Lugol’s iodine solution to 200 mL sample). Once Lugol’s is added to the sample, it requires no additional treatment prior to analysis (e.g. chilling etc.). Lugol’s iodine solution is made by mixing 20 g of KI with 200 mL of distilled water, and then dissolving 10 g of pure iodine in this solution. Glacial acetic acid (20 g) is added a few days before use. The solution must be stored in the dark in a glass bottle and remains effective for at least 12 months.

4.2.5.2 Sampling for cyanotoxin analysis

Cyanotoxin analysis will generally be required in one of the following circumstances:

• Action Level 1 status (i.e. more than 2000 cells mL-1) predominated by Microcystis aeruginosa, or when concentrations of other potentially toxic taxa (see Table 4.2) exceed 15 000 cells mL-1

• Action Level 2 status where numbers of a cyanobacterial taxa not previously recorded as toxic exceed 100 000 cells mL-1 (recommended toxicity analysis by mouse bioassay or comparative method).

Samples for toxin analysis should be collected using the 5 m integrated hose-pipe sampler (as described in Section 4.2.5) and a 2 L chilled sample sent to the laboratory for analysis (see Appendix C6 for laboratory contact details).

In Australia, and internationally, guidelines for cyanotoxins in drinking water supplies are being set based on the concentration of toxins in water (µg toxin L-1). Hence it is recommended that for those cyanobacterial species where the toxins are well known and characterised, i.e. Anabaena circinalis, Aphanizomenon ovalisporum, Cylindrospermopsis raciborskii and Microcystis aeruginosa, routine analysis of toxins should be carried out by high performance liquid chromatography (HPLC). HPLC analysis enables the exact concentration of individual toxins in cyanobacterial samples to be quantified, with toxin concentration reported either in terms of the mass of toxin per unit mass of cyanobacteria, or mass of toxin per litre of water. Mouse bioassay should only be used if taxa other than the ones listed above are suspected of producing toxin.

4.2.5.3 Sampling frequency

Monitoring class A1 and A2 storages are recommended to be sampled on a fortnightly basis, until the total blue-green algae cell count exceeds 2000 cells mL-1, after which they should be sampled weekly. Sampling can return to fortnightly after cell numbers fall below 2000 cells mL-1. Ideally, sampling frequency should be determined on a storage by storage basis using historical records of blue-green algal dynamics.

4.2.5.4 Sample analysis and reporting

4.2.5.4.1 Analysis precision

A suitably qualified laboratory with appropriate quality systems in place should conduct the sample analysis. When requesting the analysis it is important to state the minimum precision required of the analysis both in respect of the identification (i.e. genus or species level) and enumeration (provides the level of confidence in the result). The precision associated with the analysis of samples is directly related to the amount of analytical effort with respect to the laboratory equipment, counting effort and therefore time and staff expertise. Therefore, there is likely to be a higher cost associated with higher levels of precision. For A and B monitoring classes, it is recommended that the minimum taxonomic precision be at the genus level with species identification essential for potentially toxic species (see Table 4.2).

The counting precision is an estimation of the error associated with the estimation of abundance. It is defined as the ratio of the standard error to the mean (expressed as a percentage) for replicated counts and assumes a Poisson distribution of counting units in the counting chamber (Laslett et al 1988). The precision (counting error)

can be calculated from the total number of units (n) using the formula derived by Laslett et al (1998):

Counting error (± %) = 100 × π2

Therefore the more units counted, the lower the counting error. A summary of counting errors based on this formula is shown in Table 4.5 The minimum acceptable level of precision for public health monitoring is ± 30 per cent. It is also recommended that a minimum precision of ± 30 per cent be specified for potentially toxic species.

Table 4.5 Minimum counting error after Laslett et al (1998)

Total units counted Counting error (± %)

1 140

2 100

4 75

8 50

13 40

23 30

50 20

200 10

4.2.5.4.2 Sample analysis

Sample analysis is conducted by examining a subsample under a microscope and systematically identifying and counting algal units. Specialised counting chambers of a known volume are used in conjunction with a grid system to enable examination of a known area of the chamber, and corresponding volume of sample. The two most commonly used chambers are the Sedgwick–Rafter chamber and the Lund Cell. Both are used with an upright microscope. Both are similar in that they contain a fixed volume of sample; however, the Sedgwick-Rafter chamber is etched with a calibrated grid of 50 × 20 equal sized squares (1 mm²), whereas the Lund Cell is unmarked and is used in conjunction with a Whipple Grid, which is inserted in the microscope eyepiece.

There is no Australian standard for the analysis of planktonic microalgae; subsequently, laboratories will vary slightly with respect to their preferred method. An Australian ‘benchmark’ or recommended approach for the enumeration of cyanobacteria is given in Jones et al (2002), based on the Phytoplankton Methods Manual for Australian Freshwaters (Hötzel & Croome 1999). A generic method for the identification and enumeration of planktonic microalgae using a Sedgewick-Rafter chamber based on the benchmark approach is given in Appendix H11 as an example.

4.5.4.2.3 Reporting

Phytoplankton density is a concentration measure and should be reported as cells per millilitre. Algal biomass can also be reported as cell biovolume, which takes into account the contribution of species based on their relative size. Cell biovolume is measured by calculating an average volume for each species using formulae for geometrical shapes closest to the cell’s shape. The average volume for each species is then multiplied by the cell count for the species and all the products summed to gain a biovolume per sample in mm³ per litre. Advice on developing a cell biovolume method can be found in APHA (1992) and Hillebrand et al (1999). Cell biovolume measurements are recommended for the calculation of recreation hazard risk after the WHO guidelines in Table 4.3.

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4.2.6 Contingency plan framework for blue-green algae response Contingency plans provide an action framework that specifies appropriate management actions in response to blue-green algal cell level thresholds. While the Queensland Water Quality Task Force created a generic action framework in 1993 (see Table 4.6), agencies operating storages are encouraged to develop site-specific contingency plans to address their particular situation.

An action level framework is a monitoring and management action sequence that water treatment operators and storage managers can use to provide a graduated response to the onset and progress of a cyanobacterial bloom. The managerial response model presented as a ‘decision tree’ in Figure 4.11 provides for the assessment of a potentially toxic cyanobacterial bloom, with appropriate actions and responses, through three ‘threshold’ stages. The action framework is based on the Australian National Alert Level scheme, and reflects recent developments in the risk assessment and monitoring of cyanobacteria released by the WHO (Chorus & Bartram 1999).

Table 4.6 Generic contingency plan framework

Vigilance level

Threshold definition: cyanobacterial cell numbers 500–2000 cells mL-1

The vigilance level encompasses the early stages of bloom development, when cyanobacterial cells are first detected in unconcentrated lake or raw water samples.

• When vigilance level is exceeded, it is recommended to increase the sampling frequency to at least once a week, so that potentially rapid changes in cyanobacterial biomass can be monitored.

• Visual inspection for algal scums or accumulations of all water intakes and water recreation areas should be conducted on at least a weekly basis.

Action level 1

Threshold definition: cyanobacterial cell numbers > 2000 cells mL-1. Persistently high cyanobacterial cell numbers throughout the storage increasing or remaining high as per threshold definition

Action level 1 threshold (cyanobacterial cells > 2000 cells mL-1) is derived from the WHO guideline for microcystin-LR and the highest recorded microcystin content for cyanobacterial cells. The threshold level assumes that the species present is a microcystin producer, where raw water microcystin concentration could exceed the WHO guideline value of 1 µg L-1.

• If Microcystis aeruginosa is present at concentrations > 2000 cells mL-1 or other known toxin producing taxa (see Table 4.2) at > 15 000 cells mL-1, the conditions require a quantitative analysis of the concentration of cyanotoxin in the raw water supply, and an assessment of whether the water treatment processes available are effective in reducing toxin concentrations to acceptable levels. Ongoing analysis of algal toxins in the raw water is necessary if values exceed 1 µg L-

1.

• Continue routine weekly monitoring of raw and treated water to ensure adequate removal of algal cells and toxins.

• Implement use of alternative water supplies and consult health authorities if toxin concentrations in treated water exceed 1 µg L-1.

• Visual inspection of all recreation areas should be conducted prior to entering the water – bathers should avoid contact with cyanobacterial scums. See Table 4.3 for appropriate water recreational hazard status based on latest analysis results.

Action level 2

Threshold definition: cyanobacterial cell numbers > 100 000 cells mL-1.Persistently high cyanobacterial cell numbers throughout the storage increasing or remaining high as per threshold definition

The threshold for action level 2 (cyanobacterial cells > 100 000 cells mL-1) describes an established bloom with high biomass with the possibility of localised scums. Conditions in action level 2 are indicative of a significant increase in the risk of adverse health effects from the supply of water that is untreated or treated by an ineffective system or through primary contact water recreation or bathing activities.

• Maintain weekly or bi-weekly sampling (depending on the dominant cyanobacterial taxa present), including all sites and visual inspection of all water recreation areas for scum formation. Ensure warning signs indicate current recreation hazard status (see Table 4.3) or direct access to storage is restricted.

• Implement use of alternative water supplies and consult health authorities if toxin concentrations in treated water exceed 1 µg L-1.

Figure 4.11 Decision tree incorporating the model action levels framework for monitoring and management of cyanobacteria in drinking and recreational waters

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4.3 Sampling fish

4.3.1 General considerations

4.3.1.1 Permits and approvals

A general fisheries permit is required for all work that involves ‘fish’ as defined in Section 5 of the Queensland Fisheries Act 1994. Note that early life stages such as eggs, spat or spawn of fish are considered as fish under the Act. Before undertaking any work, ensure that a General Fisheries Permit is current and covers the work outlined in the methods of fish sampling planned.

Under the Queensland Nature Conservation Act 1992 (NCA), prior approval from the appropriate EHP work unit is required to conduct activities involving protected wildlife and places.

Under the Queensland Animal Care and Protection Act 2001, approval from an Animal Ethics Committee (AEC) is required for the use of animals for scientific purposes. This includes sampling of fish for scientific purposes. It is a legal obligation to receive the approval from the AEC in writing before any project can commence.

Specific procedures used to process fish will depend on the project objectives. Fish should always be handled and processed in a timely and safe manner, as specified in the animal ethics code.

All noxious fish are to be destroyed and disposed of appropriately, and not returned to the water. Consult the Department of Employment, Economic Development and Innovation (DEEDI) for the latest information on policies and legislation regarding the release of noxious fish.

4.3.1.2 Recording fish sampling data

A fish sampling data sheet should be used to record field data. An example is at Figure 4.12.

4.3.1.3 Choice of sampling method

The available methods covered in this manual are:

• fishing using drift nets

• fishing using tow nets

• fishing using small seine nets

• fishing using long seine nets

• fishing using fyke nets

• fishing using cast nets

• fishing using gill nets

• baited trap fishing

• electrofishing.

Figure 4.12 Example of a fish sampling data sheet

4.3.2 Sampling fish using drift nets

4.3.2.1 Drift net fishing method

A flow velocity of more than 0.05 m/sec is required to allow for drift nets to be effective, so nets can only be set in areas exceeding that velocity.

In flowing water, drift nets are set facing upstream in the upper water column to capture pelagic larvae where they are at their greatest density (Gilligan & Schiller 2003). Nets have a 3 m rope (with a float attached) and are tied to a tree or stake at least one metre out from the bank to ensure appropriate flow.

Net dimensions and mesh size should be determined according to project objectives. The net entrance should be covered with coarse exclusion mesh to prevent entry of debris and larger predators.

Drift nets should be set to sample a variety of habitats and velocity within the channel (e.g. edge vs mid stream). Velocity readings are taken at the entrance of each net and re-measured when the nets are checked after a set sampling period. This allows for the estimation of the volumes of water flowing through each net so that density can be estimated.

When checking, each net is taken to the bank or lifted into the boat to be processed. The tail (cod-end) is untied and the contents placed in a separate bucket half filled with water. The sides of each net should also be rinsed over the bucket to ensure that nothing small remains in the net. The cod-end is then re-tied and the net returned to its same location within the stream.

Processing involves pouring the contents of each individual bucket through a fine mesh sieve (250μm or 500μm) and then transferring the remaining animals and debris into a vial or plastic QA bag (depending on the amount of debris captured). The sample is then preserved in ethanol or methylated spirits, labelled and returned to the laboratory for processing.

4.3.2.2 Reference

Gilligan, D. & Schiller, C. (2003) Downstream transport of larval and juvenile fish in the Murray River. NSW Fisheries Final Report Series No. 50, NSW Fisheries, Narrandera.

4.3.3 Sampling fish using tow nets

4.3.3.1 Undertaking tow

Nets are attached to an aluminium frame and fixed to the front of the boat to minimise both boat and outboard

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effects on the sample (see Figure 4.13).

Figure 4.13 Boat with fixed aluminium net mount and attached nets

Depending on the project purpose, different sized mesh can be used. It is recommended that invertebrates are sampled using a 250 μm tow net, while fish larvae are sampled with a 500 μm net. Different nets can be fixed to the opposite side of the aluminium support mount.

Depending on the project purpose, tows can be stratified according to different habitat types (e.g. open water, timbered) and three replicate tows undertaken in each. Each tow should consist of a set distance and boat speed to allow for calculation of the volume of water sampled by the nets (e.g. 40 m tow, with the boat travelling at a constant speed throughout (0.75 m/s-1) (5000 L of water)). Alternatively, the tow can be timed (2, 5, 10 or 15 minutes depending on sample area and density of organisms) and the boat kept at a constant speed. Either way, GPS readings should be taken at the start and end of the tow.

4.3.3.2 Clearing nets

After the tow is completed, nets should be untied and brought back into the boat. One bucket should be filled with water to allow rinsing of any debris or animals into the base of the net. The contents of the net are then emptied into the sieve pair (i.e. coarse sieve overlying 250μm sieve). Finer material should be gently rinsed through the coarse sieve and then the contents of the 250μm sieve washed into a labelled vial for preservation. Samples will be preserved in vials using 90 per cent ethanol or methylated spirits and kept for later identification and enumeration in the laboratory.

4.3.4 Sampling fish using short seine nets Seine nets primarily sample smaller fish, and larger fish are rarely encountered.

Seine nets are most effective when used in wade-able water that is no deeper than waist height.

Seine nets only sample fish that are close to the bank at the time of sampling, so will not capture species that prefer deeper water.

Very fast moving fish swim away from the net and therefore may not be sampled effectively when using this method.

It may be difficult, if not impossible, to use a short seine net in very fast flowing water.

Two people are required with one person holding each end of the net, preferably putting a foot through a foot hole in the bottom rope to assist in keeping the bottom of the net pulled taught and on the stream bed while dragging the seine through the water in order to prevent fish from escaping. The net will snag easily, so try to avoid areas of dense logs and branches.

Position one person close to the bank and the second person out in deeper water, then drag the net briskly through the water. The seine net is dragged perpendicular to the bank along a predefined length of stream (5 metres is a practical length to use). After covering this set distance, the person in deeper water should rapidly swing around to approach the bank and both people should then quickly drag the net towards the bank, through the shallows and out of the water (see Figure 4.14). Ensure that no gaps are created between the net and the streambed to prevent trapped animals from escaping underneath. If snags are encountered, stop dragging the net, free the net from the snag, and restart seining in a different stretch of stream. Speed is required when using the net; otherwise fish will easily swim clear. The seine net should be used at least three times at each site to ensure the sampling of all the different habitats available.

It is easiest to remove captured animals when the seine net is laid on the bank. The seine net should be thoroughly rinsed after all of the fish have been released.

Figure 4.14 A seine net hauled on to the bank, enabling the removal of trapped animals

4.3.4.1 Variation to method

A variation to this method is to seine upstream heading into a riffle, instead of seining towards the bank. Each person must then lift the edges of the net out of the water, so that the fish do not escape and carry the net towards the bank. This technique may be useful for catching species that often occur near riffles.

4.3.5 Sampling fish using long seine nets

4.3.5.1 Long seine fishing method

Long seine nets have coarse mesh; therefore, smaller fish such as Hypseleotris sp. are not usually sampled.

Seine nets are most effective when used in wadable water that is no deeper than waist height. See variation for deeper water at Section 4.3.5.2.

Long seine nets only sample fish that are present close to the bank at the time of sampling. Fish that only occur in deeper parts of the river may not be encountered.

Very fast moving fish may swim away from the net, and therefore may not be sampled sufficiently when using this method.

It may be difficult, if not impossible, to use a long seine net in very fast flowing water.

It takes two people to use a long seine net, one person in the water and the other person on the bank. The person on the bank holds the top and bottom rope of one end of the net. The person in the water holds the top rope of the other end of the net and puts their foot through a foot hold in the bottom rope of the net (this will assist dragging it through the water and prevent fish from escaping under the net). This person should then wade out from the bank pulling their end of the net until all of the net has been deployed into the water (see Figure 4.24). Then this person should ‘loop’ around towards the bank until they reach the starting point. This process should be carried out reasonably quickly; otherwise, fish will easily swim clear of the net. In flowing water ensure that the loop is started by pulling the net into the current.

Seine nets are most effective in areas where snags are not present. If snags are encountered, stop dragging the net, and free the net from the snag and restart seining in a different stretch of stream.

When the loop is complete, each person then holds the top and bottom end of their end of the net and together drag the net into the shallows and out of the water. Ensure that no gaps are created between the net and the stream bed, or else fish may escape underneath. If snags are encountered, stop dragging the net, and free the net from the snag if possible; then continue dragging the net in to shore.

It is easiest to remove fish when the seine net is laid on the bank. Rinse the seine net after all of the fish have been removed.

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Figure 4.15 A long seine net being deployed from the bank

Figure 4.16 Fish being captured in the long seine net

NOTE: Long seine nets will snag easily, so ensure that the net will not be dragged around logs or branches that may be visible. If snags are unavoidable restart seining in a different stretch of stream. It is easiest to use a long seine net in slow flowing water, or in water that has stopped flowing. Backwaters may be suitable for sampling at sites with high flow.


Where the water is too deep to wade through, the technique can be varied so that a person holds one end of the long seine at the bank and the other end is looped around by boat. However, this is not ideal, as once the water is deeper than the depth of the net, fish can potentially escape by swimming underneath. It is not recommended to ‘swim’ the net out if deep water is encountered. This is a slow process and will result in a low catch as fish can easily escape. It is also potentially dangerous, as the net could wrap around the swimmer’s legs, causing injury or drowning.

4.3.6 Sampling fish using fyke nets

4.3.6.1 Fyke net fishing method

Fyke netting is a passive method of sampling and therefore may result in some species not being sampled, rare species being common but poorly sampled, or rare species that move in schools appearing more abundant than they truly are (see Figures 4.25–4.32).

Using upstream facing fine-fyke nets during a flow event can result in the net filling with or being damaged by drifting debris. These nets should be monitored to ensure that damage is not occurring. Their use may be prohibited in areas where there are large amounts of debris.

Fyke nets are set facing downstream if the aim is to capture upstream migrating fish, or set facing upstream if the aim is to capture downstream migrating fish, drifting larvae or eggs. Net mesh size and the number of replicates

should be based on the objectives of individual projects.

The nets are set at approximately 30 degrees to the bank with the wings angled out from the mouth of the net at approximately 45 degrees. The tail (cod-end) is either tied off to a tree, rock, or stake, or allowed to flow freely. A float should be placed inside the cod-end of the net and care taken to maintain an air-space so that capture of air-breathing animals (e.g. turtles) doesn’t result in mortality. Once the cod-end is tied off, wade or drive the boat into deeper water with the remaining net, spreading out the inner hoops. Note that a fyke net does have a ‘top’ and ‘bottom’, with the top having floats attached to the wings and the bottom having weights attached. Each wing should be stretched out separately (one toward the bank and one toward the deeper channel) and secured to a tree or stake. Floats on the top of the net will improve net visibility to ensure that any boats can avoid the net.

When checking the nets, pick up the first hoop. This hoop should be shaken, ensuring that any fish fall down into the next hoop. Each hoop should be shaken in turn, until all fish have been shaken down to the cod-end of the net. The tail can then be untied from its fixed position and carried to shore for sorting, or contents can be poured into a nally bin (partially filled with fresh water) in the boat.

The fyke net should be rinsed after netting has finished. Upon returning from a field trip, the nets should be thoroughly cleaned, freed of debris, hung out to dry, checked for holes and repaired if necessary before being stored.


Ethanol may be used in place of methylated spirits to preserve samples.

Figure 4.26 Setting a fyke net

Figure 4.27 A set fyke net

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Figure 4.29 A set fyke net

Figure 4.30 Opening up the tail of the fyke net to identify the captured fish

Figure 4.32 Drying fyke nets

4.3.7 Sampling fish using cast nets To start, make sure the net is free of tangles and debris, and then lay it on the ground in a horizontal line away from you (see Figure 4.33). Make a loop in the draw string of the net and place it around the wrist of your throwing arm.

Figure 4.33 Cast net

Commence measuring off approximately 1 m long loops of draw string and lay them into the palm of your hand until you get to the top of the net. Next, measure off one loop of net (for a 2 m net) and lay that too in the palm of your hand. For a net 2.4 m or larger you will need to make two loops of the net

At this point (see Figure 4.34), the palm of your hand should be holding the loops approximately level with your knee to lower thigh region. Now with your left hand, grab the bottom of the net, or leadline, and place it into the palm of your right hand (which is still holding the loops). The net is now divided equally in two, either side of your right wrist.

Figure 4.34 Cast net stage 1

With your left hand (see Figure 4.35) reach over to your right hand side (with the back of your hand facing away from you) and grab the right hand side of the lead line. Place the lead line in the cradle between your thumb and forefinger and walk your fingers along the 'inside' of the net and gather about 10 handfuls of net.

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Next, lift this gathered handful up and place it under your right hand thumb (see Figure 4.36).


Now you must gather the left hand side of the net (see Figure 4.37) by grabbing the lead line again in your left hand and holding it in the cradle between your thumb and forefinger with the back of your hand facing away from you. Gather 10 handfuls along the 'outside' of the net and hold in your left hand.


To cast, hold the net like you're holding a bullfighter's cape with your right arm fully extended and parallel to the ground. Your left arm should be parallel to your right, but slightly back and lower (see Figure 4.38). Your technique will determine the equal spread of the net.


When you throw, your feet should be firmly apart for balance, and the left foot in front of the right. The casting action is more like a flicking of your arm from the right-hand side to your left shoulder, similar to a forehand tennis shot (see Figure 4.39).


Release the net—both hands simultaneously—about a third of the way through the cast. When your net hits the water your right hand should have met your left shoulder.

Once the net sinks to the bottom, slowly pull the draw string in towards you. This will close the net and trap the specimens.

Next, carefully retrieve the net and proceed to remove your catch and place it in a container of water drawn from the immediate area.

To preserve the life of your cast net, remove any debris from the webbing and dunk it in a bucket of fresh tap water before storing it away. Avoid leaving it out in the sun for extended periods.

4.3.8 Sampling fish using gill nets The site being sampled must be first investigated to determine suitability for this type of sampling, including risk from snagging up on rocks or snags due to current. This type of sampling is not recommended in very strong flows.

It is highly recommended to use a boat for deploying gill nets. Deployment is extremely difficult without a boat, and almost impossible in deep water. Gill nets should be tied to a fixed object on the bank, and set perpendicular to the

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bank in still or slow-flowing water, or increasingly angled downstream with increasing flow velocity (see Figure 4.41).

One person should be dedicated to manoeuvring the boat into position. This person must hold a recreational boat licence and be proficient in operation of this type of craft/motor under the field conditions.

One person should be responsible for tying off the net onto a fixed structure (using a suitable knot to be able to be untied under load), as well as deploying the net. A net weight should be attached to the bank end of the leadline to ensure that the net does not move. As the net is deployed, the person steering the boat should slowly reverse the boat away from where the net was tied diagonally out into the current. The last section of net to be deployed should be weighted on the leadline with an anchor to prevent movement. A float should be attached to the end of the net to make it visible to other boaters, along with a flashing light if visibility is poor or at night.

Nets should be positioned within a contiguous stretch of river, and distributed so that all nets are fished as independently as possible.

The amount of time that the nets are deployed should be relatively similar for each site sampled. At 45 minute intervals, the nets should be each checked and fish removed by the best technique that facilitates loss of scales and slime (a knife can be used to cut mesh if needed).

The fish may be measured (length), identified to species level, and/or checked for condition (lesions, ulcers, flesh samples taken), and fish species abundance may be recorded. This information should be recorded onto an appropriate field sheet (see Figure 4.19 for an example). All native fish are to be released, unless they cannot be identified. In this case, representative specimens may be preserved in methylated spirits, or on ice, and stored in a labelled plastic vial/jar for later identification. Rinse the gill net after all of the fish have been released. To dry the net, tie each end to a structure. Check the net often, as birds and other animals could possibly become tangled in the net.

4.3.9 Baited trap fishing Traps can come in a variety of shapes and sizes, but the most common trap is the ‘funnel trap’ variety (Department of Primary Industries and Fisheries, 2009). Traps can be baited or unbaited, and often a chemical light stick is added when the trap is deployed at night. Traps should be marked in accordance with permit or state fisheries legislation. Traps are mostly set from the bank, close to snags and other stream side vegetation, but they can also be set into mid stream habitats using a boat. Traps should be deployed for a standard time, into various stream habitats (including bare banks), which helps ensure the results are comparable through time. Upon retrieval, traps are emptied into a bucket of water and captured animals are processed accordingly.

4.3.10 Sampling fish using electrofishing

4.3.10.1 Electrofishing safety

In addition to general workplace health and safety considerations, there are specific hazards associated with the

Figure 4.41 Setting a gill net that has been tied to one end to a snag

use of electrofishing equipment.

The following safety material is taken from the Australian Code of Electrofishing Practice. This code should be adhered to during any electrofishing activities.

4.3.10.1.1 All electrofishing

Rubber boots or waders, plus 1000V rated linesmen gloves must be worn by all electrofishing team members during operations.

Minimum of two persons are required per sampling party.

Staff training in the use of electrofishing equipment should be undertaken prior to sampling, with a senior person with more than 50 hours of electrofishing experience to be present.

Waders and long 1000V linesman gloves are to be worn when using electrofishing equipment.

Only fish nets with long (approximately 3 m for a boat) non-conductive handles are to be used for sampling.

Equipment should be regularly maintained.

No electrofishing should be undertaken within 50 m of another boat.

Electrofishing should not be conducted during wet weather or rough water conditions.

Polarized sunglasses should be worn when undertaking sampling.

4.3.10.1.2 Boat electrofishing

The boat driver must have a current boat licence and be sufficiently experienced with the size of boat being used and in the river or lake conditions present.

The boat must be adequately stable and have ample freeboard when fully loaded with gear, crew (and allow for fish catch).

Boat must be a minimum of 3.5 m long.

For boats under 4.0 m x 1.2 m, a crew of only two is allowed and maximum generator size is 5 kVA.

Anodes must be fixed to the bow and not capable of touching any part of the boat, and cables are to be channelled or clipped to boat sides to prevent tripping.

To reduce the risk of dip netter staff accidents, fixed or removable hand rails of at least 700 mm height must be fitted and non-skid flooring is recommended.

Non-slip flooring is required in the boat.

The drivers of electrofishing boats must use foot-operated deadman switches which must be operated simultaneously with boat netters, who must have either a one foot operated switch (several may be connected in parallel if more than one netter is used) or ‘life-line’ belt cord cut-out switches.

All lighting and ancillary electrical equipment must be extra-low voltage (less than 32VAC or less than 115VDC).

Large red DANGER warning signs must be displayed on each side of the boat.

Generators and control boxes must be fixed in position during operation.

4.3.10.1.3 Backpack electrofishing

Backpack units must meet IP 37 standards

Battery powered backpack units must use only fully sealed dry cells as a power source.

The backpack unit must incorporate a quick release harness, a deadman switch on the anode pole, and be fitted with a mercury tilt switch that cuts off power input from the battery or generator whenever the unit is tilted at more than a 45 degree angle. The tilt switch can have an automatic reset, although a manual button that can be reset by the operator is recommended. The unit must have an audible alarm when in use.

Backpack electrofishing is not recommended in water deeper than operator crotch depth.

If a backpack unit is operated from a boat, the cathode must be isolated from the boat hull. It is also recommended that two foot-pedal safety switches connected in series are incorporated.

Life jackets must be worn by backpacking crews in any dangerous situation where the water depth is greater than 0.5 m.

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4.3.10.2 Electrofishing method

Backpack electrofishing is limited to water that is not deeper than the operator’s crotch depth.

Boat electrofishing is limited to areas that are large enough for and accessible by boat.

The success of electrofishing is limited in highly turbid waters where dip netters find it difficult to see fish.

The location and number of shots will vary according to project requirements. The number of macrohabitats sampled (i.e. pool/run/riffle), and the number of microhabitats sampled within each macrohabitat (i.e. LWD/open water/smooth bank/complex bank), will be dependent on project requirements.

Whether sampling for targeted species or fish communities, shots should be as independent as possible.

Where possible, the location of each shot should be randomly selected at a site. These shots should not overlap, and adjacent shots should have a 10 m buffer between them.

The ‘power-on’ time of each shot will be project-specific; however, it must be standardised within a project. It is estimated that at least 120 seconds ‘power-on’ time should be achieved per shot.

Power settings do not need to be standardised, instead use the settings which maximise fish catch efficiency at each site.

4.3.10.2.1 Sampling for targeted specific species or complex habitats

Favoured habitats or areas of complex habitat (e.g. snags, root masses, undercut banks) can be specifically targeted during sampling. This type of sampling is aimed at maximising catch rather than providing a comparable ‘across site’ community survey. One example of applying this type of sampling is maximising the catch of golden perch for tag and release or otolith removal.

Given that time is not important to the targeted sampling approach, it is not essential to follow specific guidelines if catch per unit effort (CPUE) data are not required. However, if CPUE is to be calculated, the following guidelines should be utilised. Complex habitats should be selected and targeted, with the habitat type and total ‘power-on’ time spent in each habitat recorded on the data sheet. The time spent in individual habitats (see method below) will vary with success of fish catch and/or size of the complex habitat; however, keeping a record of total time is critical to the calculation of CPUE.

4.3.10.2.1.1 Backpack sampling

Electrofishing using a backpack unit is best for small, slower-flowing streams. A minimum of two people are needed, one to operate the electrofishing unit and one to collect specimens using a dip-net. Sampling should be conducted within at least 1 m of the selected habitat. Each shot should commence as the operator approaches the selected habitat, and continue while the operator walks around the habitat. A minimum distance of 3 m between habitat types should be employed to avoid influencing sampling at the next habitat.

For each complex habitat, the habitat type, total ‘power-on’ time and species details should be recorded.

Fish should be processed in a timely manner at the end of each complex habitat, and returned to the water.

4.3.10.2.1.2 Boat sampling

Teams will consist of a minimum of two people: one to operate the unit, and one to net specimens.

The boat should approach a complex habitat in such a way as to minimise access issues. Shots should commence while the boat approaches the habitat, continue while the boat is stationary or manoeuvring within the habitat, and while the boat exits the habitat, before the power is switched off. A minimum distance of approximately 5 m between sampled habitats should be employed to ensure that the previous shot does not influence the next sampled habitat.

For each complex habitat, the habitat type, total ‘power-on’ time and species details should be recorded. Fish should be processed in a timely manner, at regular intervals, and returned to the water.

4.3.10.2.2 Fish community sampling

Effort should be made to sample all habitats in approximate proportion to their occurrence at the site. Shots should be performed on alternate banks to cover all habitat types. Include mid-channel shots where necessary.

4.3.10.2.2.1 Backpack sampling

Electrofishing using a backpack unit is best for small, slower-flowing streams. A minimum of two people are needed, one to operate the electrofishing unit and one to collect specimens using a dip net.

The operator should attempt to thoroughly electrofish the shot area, moving in a zig-zag motion, starting from the bottom of the shot zone. The dip netter should stand downstream of the operator while dip netting fish. Nets can be set up at the upper and lower ends of a stream section to prevent movement of fish out of the sample area.

4.3.10.2.2.2 Boat sampling

Teams will consist of a minimum of two people: one to operate the unit, and one to net specimens.

Parallel runs with the boat travelling parallel to the bank can be carried out to capture mid-water and pelagic individuals.

Perpendicular runs can be carried out with the boat travelling perpendicular to the bank to capture individuals near the edge and associated habitats. At the completion of a shot, the details of captured fish should be recorded and the fish released.

4.3.10.3 Data collection

Note that when entering data into AQEIS, database field terminologies are specific. Each site requires a site number. If new site numbers are required they should only be allocated following consultation with hydrographic staff. Sample runs are multiple survey runs which are linked to a specific site on a day or within a specified date range (e.g. all shots conducted at site 123400 on 01/01/08). Samples are individual runs or shots within a sample run. AQEIS generates new sample numbers in the order of data entry but original run/shot numbers can also be retained. Specimens are the individual fish captured within a sample.

AQEIS also generates specimen numbers obtained within each sample in the order of data entry (i.e. commencing at 1 each time).

There are two EFISH sample properties templates in AQEIS, one each for boat and backpack electrofishing methods (both using Smith Root equipment). Any variables that are not collected in the field can simply be passed over when entering data. Additional variables or alternate templates can be added in by the AQEIS administrator if required. The included variables for boat fishing are shown below.

GPS Start Latitude (GDA94) Max Sample Velocity (m/s)

GPS Start Longitude (GDA94) Min Sample Velocity (m/s)

GPS End Latitude (GDA94) Av Sample Velocity (m/s)

GPS End Longitude (GDA94) Weather at Sample Time

Sample Length (from GPS) Conductivity (µs/cm)

EFISH Start Time (time on) EFISH Boat Current Mode

EFISH End Time (time off) EFISH Boat Pulses per Second

Max Sample Depth (m) EFISH Boat Voltage Range

Min Sample Depth (m) EFISH Boat Percent of Range

Av Sample Depth (m)

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The template for backpack fishing is virtually the same, but the last four variables are replaced with EFISH backpack volts, EFISH backpack frequency and EFISH backpack duty cycle.

The EFISH specimen properties template in AQEIS contains the following variables.

Fish Species

Fish Standard Length Mm

Av Depth at which fish were caught M

Otoliths Removed No by default

Gonads Preserved No by default

Stomach Contents Preserved No by default

Fin Clips Preserved No by default

Muscle Tissue Preserved No by default

Variation to method

Additional habitat properties may also be recorded. Those found in AQEIS under the EFISH habitat properties template are as follows.

Efish Habitat Type (run/pool/rifle etc) Efish Habitat % Macrophytes

Efish Habitat % O/head Canopy Cover Efish Habitat % Filamentous Algae

Efish Habitat % Mud (<0.06mm) Efish Habitat % Leaf Litter

Efish Habitat % Sand (0.06-2mm) Efish Habitat % Overhanging Veg

Efish Habitat % F Gravel (2-16mm) Efish Habitat % Emergent/Inundated Veg

Efish Habitat % C Gravel (16-32mm) Efish Habitat % Root Mass

Efish Habitat % Pebble (32-64mm) Efish Habitat % Undercut Banks

Efish Habitat % Cobble (64-128mm) Efish Habitat % LWD

Efish Habitat % Boulder (128-512mm) Efish Habitat % SWD

Efish Habitat % Bedrock (>512mm) Efish Habitat % Bare

NOTE: ‘EFISH’ is attached to the variable names to distinguish them from existing variables relevant to other projects. ‘EFISH’ can also be used as a keyword when searching the database.

Project Project Code (AQEIS) Description of variation, rationale and any project specific variables

4.3.10.4 References

Fitzroy Basin Resource Operation Plan (2004)

Ecological Monitoring and Assessment Program Pilot Study—Part 2 Research Project Indicator Methods, Department of Natural Resources, Brisbane.

NSW Fisheries (1997) Australian Code of Electrofishing Practice, NSW Fisheries Management Publication.

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4.4 References APHA & AWWA (1992) Standard methods for the examination of water and wastewater, 18th edn, prepared and published jointly by American Public Health Association, American Water Works Association, Water Environment Federation, Washington.

Chorus, I. & Bartram, J. (1999) Toxic cyanobacteria in water, E & FN Spon on behalf of the World Health Organization, London.

Department of Primary Industries and Fisheries. (2009) Recreational fishing rules and regulations for Queensland; a brief guide. pp 11.

Jones, G., Baker, P.D., Burch, M.D. & Harvey, F.L. (2002) National protocol for the monitoring of cyanobacteria and their toxins in surface waters, Draft version 5.0, ARMCANZ, National Algal Management.

Hillebrand, H., Dürselen, C.D., Kirchtel, D., Pollinger, D. & Zohary, T. (1999) Biovolume calculation for pelagic and benthic microalgae, Journal of Phycology, vol. 35, pp. 403-424.

Hötzel, G. & Croome, R. (1999) A phytoplankton methods manual for Australian freshwaters, LWRRDC Occasional Paper 22/99, Canberra.

Laslett, G.M., Clarke, R.M. & Jones, G.J. (1997) Estimating the precision of filamentous blue-green algae cell concentration from a single sample, Environmetrics, vol. 8, pp. 313-339.

NH&MRC/ARMCANZ (1996) Australian drinking water guidelines, National Health and Medical Research Council, Agriculture and Resource Management Council of Australia and New Zealand, Commonwealth of Australia.

Pilotto, L.S., Douglas, R.M., Burch, M.D., Cameron, S., Beers, M., Rouch, G.J., Robinson, P., Kirk, M., Cowie, C.T., Hardiman, S., Moore, C. & Attewell, R. (1997) Health effects of exposure to cyanobacteria (blue-green algae) during recreational water-related activities, Australian and New Zealand Journal of Public Health, vol. 21, pp. 562-566.

World Health Organization (1998) Guidelines for drinking water quality, 2nd edn, Addendum to vol. 2, Health Criteria and Other Supporting Information, World Health Organization, Geneva.

Part E Preparation of aquatic animal tissues (fish and crustaceans) for veterinary laboratory examination

5.1 Reasons for sending aquatic animal tissues for veterinary laboratory examination There are several reasons why aquatic animals associated with a fish kill, or other environmental health or aquaculture production problems, should be submitted to a diagnostic laboratory for examination:

• The clinical signs and gross lesions in aquatic animals may not indicate the particular disease, i.e. they are non-specific signs, and laboratory examinations are required to determine the actual cause of a disease and to then provide the diagnosis.

• A laboratory examination and laboratory testing is necessary in order to make a definitive or confirmed diagnosis in diseases where a provisional diagnosis has been made on history and clinical signs.

• It is not uncommon to have more than one agent or factor affecting aquatic animals at one time, e.g. a bacterium, a fungus and handling damage to the skin. Laboratory examinations aid the identification of primary and secondary aetiologies.

• An accurate diagnosis is often a complex process requiring the skills of several scientists who are available in the diagnostic laboratory system.

• Appropriate and responsible treatment and control of disease in aquatic animals depends on an accurate diagnosis. The negligent or unnecessary use of chemicals and antibiotics may result in chemical residues in aquaculture products for human consumption, unnecessary production costs, antibiotic resistant strains of bacteria and environmental damage.

• A wide range of environmental and/or husbandry factors influence the expression of disease, and diagnostic laboratory examinations may be required to determine the significance of infectious agents in a disease.

Many diseases in aquatic animals are a result of a complicated interaction between the animal, the environment and a pathogen or disease agent (bacteria, fungi, parasite, virus and toxins). Environmental factors (temperature, salinity, etc.) act as triggers in these situations allowing a pathogen (disease producing organism) to cause disease where it might normally be harmless (see Figure 5.1).

Figure 5.1 Disease results from the interaction of animals with environmental factors and disease organisms

Environmental factors that are involved in contributing to disease outbreaks—for example, low pH, low dissolved oxygen concentrations, pesticides and high suspended solid concentrations—can also directly affect the health of aquatic animals. This type of disease is often described as non-infectious disease.

As a consequence, it is important to determine the role and significance of both the environmental factors and the pathogens. The ‘history’ of the disease outbreak, site investigations and laboratory examinations are all necessary to really understand how the aquatic animals become sick.

The best way to control or prevent a disease may be the manipulation of the environmental factor(s) that make the aquatic animal susceptible to an infectious disease.

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In any disease investigation, the following are needed:

• a history or details of the events leading up to and associated with the disease outbreak. This should include water quality, source of animal, stocking density, distribution of mortalities by pond or age group, feed type, and so on

• samples of both ‘sick’ and ‘healthy’ animals

• other samples may also be necessary, e.g. water, algae and feed.

5.2 Collecting finfish specimens for diagnostic laboratory examination The best samples to send to the laboratory are live finfish. Live specimens are best as they enable the pathologist to see the clinical signs, prepare gill and skin smears and are suitable for all laboratory tests (histology, bacteriology, parasitology, virology, biochemistry and toxicology).

Because finfish are in an aquatic environment and carry bacteria on the gills and in their gut, there is a rapid breakdown of tissue and decomposition after death. In most cases aquatic animals that are found dead may be of little value for veterinary laboratory examination.

5.2.1 Sampling live finfish Select a minimum of six live, sick fish with clinical signs typical of the problem. Select six normal fish for comparison. A sample from each pond, cage, tank or aquarium, or each age class or species affected in the disease outbreak should be provided. Place each group in a separate and labelled container.

Transport the live specimens to the laboratory in oxygen filled plastic bags or aerated transport tanks.

5.2.2 Sampling and preparing fixed finfish specimens Use only live finfish for fixation (fish that are found already dead are of little value for veterinary diagnosis).

Painlessly kill the finfish in an anaesthetic solution or by cutting the spinal cord behind the head.

A complete external and internal examination of the finfish should be done and the findings recorded on the specimen advice sheet to the laboratory.

Microscopic examination of wet preparations made from skin and gill smear for external parasites is essential as these parasites will leave the fish after it is placed in the fixative.

• Larval finfish: preserve 25 to 50 individuals intact

• Small finfish (less than 4 cm in length): cut open the abdominal cavity, move the swim bladder and visceral mass away from the caudal kidney, and place in fixative

• Large finfish (more than 4 cm in length): the individual organs should be removed, sectioned and placed in fixative. Tissues to sample include the gills, heart, liver, spleen, head kidney, caudal kidney, digestive tract (intestine), eye, skin, muscle, brain, swim bladder and any other organ or tissue with lesions. Tissue pieces should be about 15 x 15 mm and no thicker than 8 mm.

Place the tissues in chilled fixative if possible. There must be at least 10 times more fixative volume than tissue volume. After 24 to 48 hours, the fixed tissues can be wrapped in fixative-moistened paper towels, placed in two sealed plastic bags and sent to the laboratory.

The fixatives used include 10 per cent formal saline (for marine specimens), 10 per cent buffered neutral formalin (for fresh and brackish water specimens), and Bouin's solution (has the advantage of rapid preservation).

NOTE: Fixed specimens are only suitable for pathology and histopathology (not chemical analysis).

5.2.3 Freshly killed finfish on ice Fish on ice are only of use for bacteriology, virology and toxicology. If you submit freshly killed fish on ice, they must be accompanied by a submission of fixed specimens.

Finfish are painlessly killed with an anaesthetic, wrapped and sealed in a plastic bag, and placed in an excess of crushed ice.

The delay between sampling the finfish and arrival at the laboratory should be no longer than 24 hours.

NOTE: Freshly killed finfish on ice are only suitable for bacteriology and internal parasite examination, although they can be used for virology, toxicology and histopathology if necessary.

5.2.4 Frozen finfish These specimens are only suitable for toxicology and virology. Specific suspect toxin(s) (e.g. pesticides) need to be nominated before laboratory analysis will be done.

5.2.5 Fixatives and anaesthetics Have plenty of fixative on hand, as you need at least 10 times the volume of fixative for each volume of tissue or finfish.

5.2.5.1 Fixatives

10 per cent formal saline (for marine specimens):

• formalin (37–40 per cent w/v formaldehyde solution)—100 mL

• saline solution (0.73 per cent, 73 gm salt per 10 L water)—900 mL

(Saline solution may be replaced with seawater for marine finfish.)

NOTE: formalin is a skin and respiratory irritant, and care must be taken when using it.

10 per cent buffered neutral formalin (for fresh and brackish water specimens):


• tap or distilled water—900 mL

• disodium hydrogen phosphate Na2HP04.2H20—6.5 grams

• sodium dihydrogen phosphate NaH2P04.2H20—4.5 grams

NOTE: formalin is a skin and respiratory irritant, and care must be taken when using it.

Bouin's solution:

• saturated aqueous picric acid—750 mL

• formalin (37-40% w/v formaldehyde solution)—250 mL

• glacial acetic acid—50 mL

NOTE: Specimens must be transferred from Bouin's solution to 70 per cent ethanol (ethyl alcohol) after 24 hours. Store tissues in 70 per cent ethanol.

NOTE: Dry picric acid is explosive. Use extreme care with storage and handling.

Davidson's solution:

• glacial acetic acid—115 mL


• 95 per cent ethyl alcohol (ethanol)—330 mL

• tap or distilled water—335 mL

NOTE: Once specimens have been fixed in Davidson's solution for 24 to 72 hours they must be transferred to 50–70 per cent ethanol. The fixed tissue is then stored in 50–70 per cent ethanol.

5.2.5.2 Anaesthetics for euthanasia

MS 222 (Tricaine methanesu1phonate; ethyl m-aminobenzoate; 3-aminobenzoic acid ethyl ester) 1 gram per litre of water. This concentration is lethal in 5 to 10 minutes.

Benzocaine (Ethyl p-aminobenzoate; 4-aminobenzoic acid ethyl ester):

• Benzocaine is not soluble in water and a concentrated stock solution should be made up.

• Dissolve 50 grams of benzocaine in 500 mL of 95 per cent ethanol (100g/L). If stored in a dark bottle the stock solution will keep for at least a year.

NOTE: 10 mL of benzocaine stock solution per litre of water (19/L) is lethal in 5 to 10 minutes.

Benzocaine is less expensive to use than MS 222.

AQUI-S (2-methoxy-4-propenyl phenol)

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A two stage process is suggested when using AQUI-S for anaesthesia and humane death:

• Stage 1: use 25 to 30 ppm to induce heavy sedation (i.e. no gill movement, no cough reflex on forced extension of the operculum)

• Stage 2: humanely kill the fish by pithing.

NOTE: All chemical anaesthetics must be prescribed by a veterinarian for use in food fish.

5.2.6 Basic anatomy of finfish Actual details of finfish anatomy may vary from species to species. The generalised diagrams and the photographs given below indicate the general location of organs (see Figures 5.2 and 5.3).

Figure 5.2 External anatomy of a finfish.

Figure 5.3 Internal anatomy of a finfish.

5.2.7 Gill and skin smears and wet mounts Microscopic examination of mucus and surface cells from skin and gill scrapings is the best way to find external parasites. External parasites will fall off or leave the host during transport, after death or after fixation, so it is important that only freshly dead finfish are used for preparation of smears. Smears should be examined before specimen fixation.

5.2.7.1 Method for preparing gill smears and wet mounts for finfish

• Painlessly kill the fish by cutting the spinal cord behind the head.

• Remove a gill arch, place on a clean glass microscope slide and gently scrape mucus off the gill filaments with a clean scalpel blade onto the clean glass microscope slide (see Figure 5.4 (3)).

• Gently scrape the skin, particularly areas at the base of pectoral and pelvic fins, with a clean scalpel blade and place the mucus on a clean glass microscope slide. Scales should not be removed when preparing skin smears (scrape in the direction of the scales) (Figure 5.4 (4)).

• Add a drop of water that the fish were swimming in and gently mix (see Figure 5.5).

• Cover with a cover slip (Figure 5.5). Try and avoid trapping air bubbles.

• Examine under a microscope using a low-power objective (either 10x or 20x). Examine smears as soon as possible as the movement of living parasites greatly aids detection. Do not let the wet preparation smear dry out, as the parasites will die quickly.

Figure 5.4 Preparing a skin and gill smear

5.2.7.2 Important points concerning smears and wet mounts

Any cysts or nodular structures found during external or internal examination can be removed, squashed or teased apart, wet mounted and examined in the same way that gill/skin smears are prepared.

At times it can be useful to cut off entire gill filaments and wet mount them for microscopic examination. Again, external parasites will be apparent and some indication of gill damage is seen by studying the filament and lamellar structure. In the case of bacterial gill disease, bacteria and areas of hyperplasia and necrosis will be seen.

Figure 5.5 Preparing a skin and gill smear

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5.2.8 Finfish dissection The basic method of finfish dissection involves three or four cuts (see Figure 5.6):

1. Lay the fish flat on one side with the dorsal fin facing away from you.

2. Lift the operculum (gill cover) and cut it off being careful not to damage the gills (cut 1).

3. Make a small cut with scissors or a scalpel blade just in front of the anus (vent) to open the abdominal cavity. Avoid cutting the lower intestine or damaging any abdominal organs when you do this.

4. With blunt ended scissors, cut along the ventral midline forward from the small cut to between the jaw (cut 2).

5. Remove the flap of skin covering the abdominal cavity by cutting from the small cut upwards and forwards (cut 3). The heart and all the abdominal organs should be exposed for examination and removal (see Figures 5.7 and 5.8).

6. If the brain is to be examined, remove the top of the head with several shallow slices using a scalpel blade, cutting from just behind the eyes towards the front (cut 4). Alternatively, the whole head can be removed; place the cut end down and make the slice downwards towards the dissection board. This can be done if the fish is young and the skull is not too hard. In older fish, bone cutters or saws will need to be used.

Figure 5.6 Partial dissection of a finfish using four cuts

Figure 5.7 The internal anatomy of a dissected finfish. AF = abdominal fat, G = gills, H = heart, HK = head kidney, L = liver, PC = pyloric caecae, S = stomach, SB = swim bladder

5.3 Collecting crustacean specimens for diagnostic laboratory examination The crustacea include prawns, crayfish and crabs.

Prawns, crabs and crayfish decompose very rapidly after death. Only live or freshly fixed specimens are useful for veterinary laboratory examinations. The hepatopancreas (digestive organ), which is important in the diagnosis of many diseases, decomposes extremely quickly and special care needs to be taken to ensure it is preserved correctly.

5.3.1 Sampling live crustacea The best samples to send to the laboratory are live samples. Live specimens are preferred as they enable the pathologist to see the clinical signs and prepare gill wet mounts, and are suitable for all laboratory tests (histology, bacteriology, parasitology and toxicology). Transport the live specimens to the laboratory in oxygen filled plastic bags or aerated transport tanks (see section H).

Select a minimum of six live, sick animals with clinical signs typical of the problem. Select six normal animals for comparison. A sample from each pond, cage, tank or aquarium, or each age class or species affected in the disease outbreak should be provided. Place each group in a separate and labelled container.

For larval and early post-larval crustaceans (less than 10 mm) or crab-1 etc, sample 50 larvae or post-larvae from each rearing or nursery tank, or age class or species affected by disease or mortality.

For juveniles (more than 10 mm long) and adult prawns or crabs, select a minimum of six live, sick animals with clinical signs typical of the problem together with six normal animals for comparison. A sample from each pond, pool, or tank, or age class or species affected in the disease outbreak should be provided.

5.3.2 Sampling fixed specimens If it is too difficult to transport the specimens alive, the only alternative is to send fixed specimens. Fixed specimens are suitable for pathology and histopathology.

Use only five crustacea for fixation.

Juveniles and adults can be anaesthetised by chilling in a freezer. Do not freeze them.

A microscopic examination of larvae or post-larvae, and external and internal examinations of juvenile and adult crustacean, should be done and the findings recorded on a note to the laboratory.

Figure 5.8 The internal anatomy of a dissected finfish. This photograph shows the location of the caudal kidney (CK) which is exposed when the swim bladder is deflected ventrally. The caudal kidney lies against the vertebral column and runs the length of the abdominal cavity, L = liver. SB = swim bladder

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Examination of wet preparations of gill filaments and appendages for attached organisms or abnormalities should be done if necessary before fixation.

5.3.3 Basic anatomy of crustacea Before embarking with specimen fixation, familiarise yourself with the general anatomy of the species you are dealing with (see Figures 5.9 – 5.12).

Figure 5.10 Internal anatomy of a prawn. FC = foregut chambers, Ha = haemocoel, H = heart, HG = hind gut, MG = mid gut, MGG = mid gut gland, O = ovary, VNC = ventral nerve cord.

Figure 5.9 External anatomy of a prawn. A=antenna, AB=abdominal segment, AC = adostral carina, AF = antenna flagellum, AS = antenna scale, E = eyestalk, HS = hepatic spine, P = pereiopods, P1 = pleopods, R = rostrum, SAS = sixth abdominal segment, T = telson, TM = third maxilliped, U = uropod

Figure 5.11 External anatomy of a crab. A = antenna, AB = abdomen, DG = digestive gland, E = eye, SL = swimming leg, WL = walking leg

5.3.4 Sampling and preparing fixed crustacean specimens

5.3.4.1 Dissecting prawns and crayfish

The method of prawn and crayfish dissection varies according to the size of the animal:

• Larvae and post-larvae less than 10 mm (1 cm) in length (or crab-1):

o Catch 25 to 50 individuals with a plankton mesh net and place whole into a bottle of fixative solution, either 10 per cent formalin or Davidson’s solution.

• Prawns or crayfish 10 mm to 30 mm in length:

o Cut between the thorax (head) and abdomen (tail) and place in fixative.

• Crabs 10 mm to 30 mm:

o Cut open the top carapace (shell) of the crab and place whole animal in fixative.

• Prawns or crayfish greater than 30 mm in length:

o Draw up Davidson's solution into a syringe. Attach a needle (e.g. 21 gauge). Hold prawn firmly with head facing away from you. Inject 0.5 to 2.0 mL of fixative (depending on the size of the prawn) at a 45 degree angle into the head, aiming for the hepatopancreas.

o Reposition the needle so it is pointing towards the tail and inject 1.0 to 2.0 mL of Davidson's solution into the abdominal muscle.

o Remove the thorax (head) from the abdomen (tail) by slicing between the head and tail using scissors.

o Cut along the dorsal surface (top) of the head, just underneath the carapace (shell) from the end of the head carapace right up to the rostrum. Take care not to damage the heart or stomach which lie directly beneath the shell of the head.

o Turn the prawn over and cut along the ventral (bottom) side along the midline thoracic carapace between the walking legs.

o Cut out a segment of abdomen, the thickness of one abdominal segment and place into the bottle of fixative (see Figure 5.13).

NOTE: Davidson's solution is a harsh chemical mixture and precautions should be taken to avoid skin and eye contact.

Figure 5.12 Internal anatomy of a crab. AG= antennal gland, DG = digestive gland, E = eye, H = heart, HG = hind gut, M = mid gut, O = oesophagus, S = stomach, SG = supraoesophageal ganglion, TG = thoracic gland, VNC = ventral nerve cord

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5.3.4.2 Dissecting crabs

The method of crab dissection is as follows:

• Make two longitudinal cuts with a small hacksaw from the posterior (back) edge of the carapace (shell) adjacent to the swimming legs to the anterior (front) edge of the carapace just lateral to the eye stalks (see Figure 5.14).

• Cut across the carapace from right to left swimming legs, joining the two longitudinal cuts.

• Remove the piece of dorsal carapace. When lifting the carapace, carefully dissect away muscles and tissue attachments on the inside of the carapace.

• Twist off one lateral edge of the carapace to expose the gills on that side. Remove the digestive organ contained in this lateral carapace for fixation.

• The heart and thoracic organs should be exposed for examination and removal. The thoracic ganglion and antennal gland are only visible after the stomach, heart and digestive gland are removed.

Figure 5.13 Steps in the dissection of a prawn or crayfish

5.3.4.3 Important points concerning fixation of crustaceans

The fixative of choice for crustaceans is Davidson’s solution.

If Davidson's solution is not available, 10 per cent formalin (formal saline or buffered neutral formalin) (see section 3.5.4) can be used as a poor alternative (do not transfer to ethanol).

There must be at least 10 times more fixative volume than tissue volume. After 24 to 72 hours the fixed tissues should be removed from the fixative solution, rinsed in tap water and transferred for storage in 50–70 per cent ethanol.

If possible place the tissues in chilled fixative as it will increase the time available for the chemicals to penetrate the cells, prior to decomposition.

Fixed specimens can be sent to the laboratory by wrapping in paper towels moistened with 50–70 per cent ethanol and sealing in two plastic bags.

Frozen crustaceans are only suitable for toxicology or virology. The history must indicate toxicoses and specific suspect toxin(s) must be nominated before laboratory analysis will be done.

5.3.5 Gill, appendage and larval wet mount preparations Microscopic examination of gill clips and the tips or appendage outgrowths is the best way to detect ectocommensals, ectoparasites and excessive debris. Larval crustacea are small and many lesions and tissue changes can be missed when they are examined histologically. Gut contents, heart and gut activity, bacteria, fungi, protozoans and lesions are more easily assessed when larvae are examined microscopically in wet mount preparations.

5.3.5.1 Method for preparing gill smears and wet mounts for crustaceans

Remove the gill cover with scissors, cut off a gill filament and place on a clean glass microscope slide (see Figure 5.15). Remove appendage tips or outgrowths and place on a glass slide. Collect a group of larvae by filtration or pipette and place on a glass slide.

Add a drop of water (water that the crustacean was swimming in) or saline solution on to the glass slide.

Figure 5.14 Steps to the dissection of a crab.

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Cover with a cover slip and avoid trapping air bubbles.

Examine under a microscope using a low-power objective (either 10x or 20x). Do not let the wet preparation dry out (add more water with a pipette), or the crustacean larvae and protozoa will die quickly.

Examination of larvae should cover 10 areas: eye surface, antennae, rostrum, gut content, midgut movement and size, gill area, walking legs (pereiopods), swimming legs (pleopods), tail (telson and uropods), and general pigmentation (see Figure 5.16).

5.3.6 Sampling and preparing molluscs Oysters and clams decompose very rapidly after death. Only live or freshly fixed specimens are used for diagnostic laboratory examination (see Figures 5.17–5.19).

5.3.6.1 Live molluscs

• Larvae and spat (or post-settlement stages):

o Sample 50 larvae or spat from each rearing or nursery tank/pool/pond or age class or species affected by disease or mortality.

• Juveniles and adults:

o Select a minimum of six live, sick animals with clinical signs typical of the problem together with six normal animals for comparison. A sample from each raft, rack area or tank, or age class or species affected in the disease outbreak should be provided.

• Transport the live specimens to the laboratory in oxygen filled plastic bags (larvae) or wrapped in damp paper towels or cloth in an insulated container.

Figure 5.16 Points of a prawn larvae to examine

Figure 5.15 Preparing a gill mount from a prawn

NOTE: Live specimens are preferred as they enable the pathologist to see the gross clinical signs and prepare gill and wet mounts, and are suitable for all laboratory tests (histology, parasitology, bacteriology and toxicology).

5.3.6.2 Fixed mollusc specimens

• Select only five molluscs for fixation.

• Juveniles and adults: these can be anaesthetised by chilling or by placing them in magnesium chloride or propylene phenoxetol solution.

• A microscopic examination of larval molluscs should be done and the findings recorded on a note to the laboratory.

• Larvae and post-settlement stages less than 10 mm shell length: preserve 25 to 50 individuals intact.

• Oysters greater than 10 mm shell length: cut open the two valves, remove the soft organs from the shell and place in fixative. Cut the visceral mass into pieces no thicker than 8 mm if necessary, before fixation.

• Clams up to 10 cm shell length: remove the two valves by cutting the mantle and adductor muscle attachments, and place in fixative.

• Clams greater than 10 cm shell length: remove the valves, dissect out individual organs. Tissues to take include the gills, digestive gland, mantle, kidney, muscle, heart, gonad and any other organ or tissue with lesions.

• Pearl oysters up to 10 cm shell length: cut open the two valves, and remove and preserve all the soft organs as a group. A shallow cut may be made on one side of the organs to allow the fixative to enter more rapidly.

• Pearl oysters greater than 10 cm shell length: cut the oyster in half by making a parallel cut between the two shell valves. Remove the two halves of oyster tissue and place in fixative. If there is too much tissue, place each half in a separate container which is labelled so that both halves can be identified to the one pearl oyster.

• If possible place the tissues in chilled fixative as this will slow decomposition before the chemicals have a chance to penetrate the mollusc cells.

• Fixed specimens can then be sent to the laboratory by wrapping in paper towels moistened with fixative or 50–70 per cent ethanol (whichever is appropriate) and sealing in two plastic bags.

• The fixatives used include 10 per cent formal saline and Davidson's solution.

NOTE: Fixed specimens are only suitable for pathology and histopathology.

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Figure 5.17 Internal anatomy of a pearl oyster. A = adductor muscle, D = digestive gland, F = foot, G = gills, H = heart, I = intestine, M = mantle, S = stomach

Figure 5.19 Internal anatomy of a clam. A = adductor muscle, B = byssus, C = ctenidia, D = digestive/reproductive organs, F/BO = foot/byssal retractor muscle, H = heart, K = kidney, S = shell valve, SM = siphonal mantle

Figure 5.18 Internal anatomy of an edible oyster. A = anus, AM = adductor muscle, DG = digestive gland, G = gill, H = hinge, M = mouth, Ma = mantle, P = palps, S = stomach

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Part F Monitoring mangrove forest health NOTE: In Queensland, mangrove and other marine vegetation is protected under the Fisheries Act 1994. Permits are required from Queensland Primary Industries and Fisheries (QPIF, under the Department of Employment, Economic Development and Innovation) to remove, destroy or damage a marine plant; or cause a marine plant to be removed, destroyed or damaged. Therefore, persons monitoring in this environment and requiring the removal of any mangrove matter from the field should contact their local office of the QPIF for further information.

6.1 Mangrove litter trapping

6.1.1 Introduction—what is litter and how is it related to mangrove forest health? Mangrove litter production is the shedding of vegetative and reproductive structures of mangroves (e.g. leaves and seeds). This may be caused by natural growth cycles, age, stress and/or mechanical factors, such as wind. This litter fall, part of the net primary productivity of a mangrove system, is the basis of detritus food chains.

The rate of litter production can indicate the health of a mangrove community. A healthy system will produce a high and/or stable monthly volume as older leaves are shed and replaced with new ones. Declining production may indicate that a community is under stress. The amount of reproductive material can also be used as a measure of health, as reproduction effort can be considerably reduced if plants are stressed.

The ratio of fallen leaves to stipules of Rhizophora spp. can also be used as an indicator of system health. Every developing leaf is enclosed in a stipule, which is shed when the leaf matures. Thus, in a healthy community, the ratio of leaves to stipules in the litter should be close to 1:1. If there are more leaves than stipules, this may indicate that the plant is shedding leaves due to stress.

A significant decline in leaf and flower production outside seasonal variations may indicate that a mangrove community is under stress.

6.1.2 Why monitor litter productivity? The main reasons for monitoring litter productivity are to:

• gain an understanding of the baseline litter productivity of a mangrove community

• indicate system health—litter fall is a useful indicator of this as mangrove communities under stress are likely to be less productive, resulting in less litter production over time. Alternatively, communities under stress produce a large amount of litter over a short time as the plants shed leaves

• look at unseasonably low litter production due to poor growth resulting in less detritus, and this may adversely affect faunal communities.

6.1.3 Method summary Leaf litter traps are installed in a mangrove community. Litter is collected monthly, sorted into different categories (leaves, twig, bark, flowers and propagules), oven dried, and weighed. The dry weight is a measure of the productivity of the community. Problems to be aware of include: significant disturbances (e.g. cyclones) may damage sites and make it necessary to begin collecting data again for a time series; traps can be interfered with by others; and climatic and biological differences in Australian mangroves make it difficult to compare data from different locations.

If a storm has damaged a site and stripped many of the leaves, litter should not be collected. Instead, traps should be emptied and checked for damage. Monitoring can recommence, but a new starting date will have to be noted. As leaf loss is normally balanced with leaf production, a 1:1 ratio is the most useful indicator of mangrove health.

Leaf litter traps can be constructed from PVC pipe and plastic mesh as illustrated in Figure 6.1.

6.1.4 Site selection Survey the mangrove community and identify the different forest types or zones that occur. Depending on the objectives of your program, select a homogenous forest type to monitor. Within this forest, put litter traps at 5 m intervals along a transect running parallel to the tidal gradient. Alternatively, put them at random in large quadrats (e.g. 10 m x 10 m, 20 m x 20 m, etc.).

6.1.5 Installing litter traps Install litter traps by attaching nylon cord to each corner of the trap and hanging them evenly from mangrove branches. Ensure that the bottom of the trap or chute is above the high tide mark (see Figure 6.2).

6.1.6 Emptying the traps Traps should be emptied every month (every two weeks if measuring Avicennia marina) to prevent leaf decay and to determine monthly trends. To empty a trap, remove any large sticks and put them in a plastic bag marked with the trap number. Put the bag under the chute, untie the chute and empty the trap contents. Re-tie the chute securely and proceed to the next trap.

6.1.7 Sorting trap contents On returning from the field, sort contents into leaves, flowers, bark (include wood), seeds, other and, if monitoring in a Rhizophora forest, stipules. Do not mix the litter contents from different traps. Count the number of leaves and stipules and record the result. Place the sorted contents into smaller, labelled paper bags, and then put the datasheet and smaller bags in the large labelled plastic bag for transport to the drying ovens.

Note: Leaf litter contents can be kept in a refrigerator for up to a week before being dried.

Figure 6.1 How to make a litter trap

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6.1.8 Dry and weigh trap contents • Dry the labelled paper bags in a drying oven at 70ºC for 72 hours.

• Using laboratory scales, weigh the contents from each category in each trap separately. Record the results (in grams, to three decimal places). If there is insect damage, leaf dry weight is likely to be low, biasing estimation of leaf productivity. Therefore, leaf loss needs to be quantified by sorting leaves from each trap into the closest matching category as detailed in Table 6.1 and Figures 6.2 and 6.3.

For example, if a leaf is about 40 per cent intact, put it in the '50% remaining' category; classify a leaf with 90 per cent remaining as a 'full leaf'. Weigh material in each percentage loss category, and correct for damage using the formulas provided. The sum of the corrected weights of leaves in all categories is the weight of leaves for the trap. Record this result on the datasheet for each trap.

Table 6.1 Percentage loss categories of mangrove leaves

Category of

leaf loss Measured weight Correction factor Corrected weight

Full leaf None (as measured)

75% remaining Multiply by 1.333

50% remaining Multiply by 2

25% remaining Multiply by 4

Total =

Figure 6.2 A litter trap in a mangrove forest

Figure 6.3 Leaf percentage loss

6.1.9 Data interpretation Data is interpreted as mean dry weight of litter fall per square metre, per month, expressed as g/m2/month. Ranges and standard deviations between traps should also be calculated.

Mangrove communities exhibit strong seasonal, annual and temporal variations in litter production, with peak fall occurring in summer in most locations, and varying with climatic conditions from year to year. It will also vary between regions due to different climate, rainfall, salinity and nutrient availability. For example, mangroves in the wet tropics are likely to produce more litter than those in more temperate climates. As different species also produce litter at varying rates, it is not usually appropriate to compare results between different regions and species.

Significant decline in leaf litter fall or reproductive effort in a particular mangrove community may indicate that it is under stress, so seek advice from relevant experts if such trends occur. If soil salinity is also being monitored, check data to see if unusual levels (high ones, in particular) have been recorded.

If comparing the stipules to leaves ratio in communities of Rhizophora spp., the ratio should normally be close to 1:1. Trends showing a higher ratio of leaves to stipules may indicate that the plants are shedding leaves, indicating possible stress.

For analysis of trends, data should be collected for at least three years, as some species of mangroves produce propagules every 2–3 years only.

6.1.10 References and further reading Bunt, J.S. (1995) Continental scale patterns in mangrove litter fall, Proceedings of the Asia Pacific Symposium on Mangrove Ecosystems, 1–3, pp. 135–40.

Duke, N.C., Bunt, J.S. & Williams, W.T. (1981) Mangrove litter fall in north-eastern Australia, 1. Annual totals by component in selected species, Australian Journal of Botany, vol. 29, pp. 547–53.

Snedaker, S.C. & Snedaker, J. G. (1984) The Mangrove Ecosystem, Research Methods, UNESCO, Paris.

6.2 Seedling regeneration

6.2.1 Introduction—why monitor mangrove seedlings? Due to their intolerance to shade, seedlings of most mangrove species are absent, or in low densities, under a mature forest canopy. However, the death of mature trees leaves a gap in the canopy, allowing increased light to reach the forest floor, and triggering the establishment of seedlings. These rapidly colonise the light gap, beginning the process of regeneration and eventually refilling it. These seedlings can be more susceptible to environmental stress than mature mangroves are. A heavy deposition of sediment from an event or major changes to site hydrology can induce stress, resulting in decreased growth rates and increased mortality.

Light gap regeneration rate can be an indicator of mangrove system health. Seedlings respond to environmental changes more rapidly than mature stands do, and monitoring can produce useful data in as little as three months, which is the normal monitoring frequency.

Long-term hydrological changes, such as sediment deposition or greater tidal and freshwater influence, may have occurred at the site over the past 10–30 years, resulting in colonisation by a different species from the original. For example, long-term sediment deposition may raise the elevation of a site, resulting in conditions more favourable for species normally found higher up the tidal gradient. Other hydrological changes, resulting in more tidal or freshwater influence, could create conditions favouring species normally found closer to the seaward or landward

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margin. Identifying what species of mangrove is replacing the previous forest can indicate whether climatic or hydrological changes have occurred.

6.2.2 Method summary In a light gap or a recovering mangrove forest, a belt transect containing a minimum of 25 mangrove seedlings is established. The height and stem diameter of each seedling within the transect is monitored every three months and is used to calculate the approximate trunk volume of the seedling.

6.2.3 Site selection When selecting a site, use recent aerial photographs or local knowledge to find canopy or light gaps in a mangrove forest, and confirm that there are seedlings present by ground truthing. At least four light gaps from one homogenous mangrove community are required for data interpretation.

6.2.4 Establishing a belt transect To establish a belt transect running north-south through the middle of a light gap:

• Walk to the middle of the gap and take bearings directly to the north and south.

• Mark with stakes the spots where the north-south bearings intercept the boundary of the gap. These will be the start and end points of the transect.

• Starting at the northern point, lay out the 50 m tape measure through the middle of the gap, to the southern boundary (see Figure 6.4)

• Record the length of the transect on the datasheet.

The transect needs to be wide enough to include at least 25 seedlings within its boundaries. Start with one that is 2 m wide (1 m either side of the tape measure), and count the number of seedlings. Increase or decrease the distance until a suitable width, containing an adequate number of seedlings, is established. Record this width on the datasheet.

If seedlings are extremely dense, establish four quadrats, each with 25 or fewer seedlings at regular intervals along the transect. The seedling density determines the size of the quadrats, but all must be equal. If seedlings are sparse (less than 50 in the entire gap), or if the light gap is small, sample the entire gap.

Note: This method is a general approach to establishing a monitoring plot that will compensate for uneven light distribution in most instances. However, a belt transect may not always be suitable, and the dimensions of a suitable monitoring plot should be established case by case.

Figure 6.4 Belt transects can be established through a light gap running north to south

6.2.5 Tag seedlings Using flagging tape, tag each seedling along the transect with a unique number. Beside the corresponding number on the datasheet, record the distance of each seedling along the transect and its position (distance to the left or right) from the middle of the transect (e.g. 12 m, left 0.7 m). This allows for easy location of the seedlings during

repeat surveys.

6.2.6 Measure height Using the measuring stick, record the height of the seedling by measuring from the ground to the base of the uppermost apical shoot. If the seedling is growing from a propagule, take the measurement from just above (see Figure 6.5). Record the result on the datasheet.

6.2.7 Measure stem diameter and stem density Using plastic calipers, measure the stem diameter of the seedling at the base of the cotyledon, which is just above the original propagule. If there is no propagule, take the measurement at the base of the stem, just above the swelling.

Density (number of stems per m2) is also recorded. As the light gap matures, density is likely to decrease naturally, as many of the seedlings die due to competition.

.

6.2.8 Count the leaves Count the number of leaves on each seedling. If there are more than 25, record the result as more than 25 leaves.

Leaf counts provide an indicator of seedling progress in the early stages of development. However, after 25 leaves have established this becomes very time consuming.

6.2.9 Record sediment type, pH and salinity Collect a sample of the substrate at the middle of the transect and rub it between your fingers. Record the sediment type (silt, fine sand, sand, coarse sand, or gravel) based on its feel, listing the dominant type first (i.e. record a silty sand as sand/silt).

It is also advisable to record the salinity and pH of the sediment within the transect.

6.2.10 Draw a mud map of the site Draw a mud map of the gap showing its dimensions, an arrow representing north, the position of the transect, and the surrounding forest type.

6.2.11 Measurements needed during re-survey When the site is re-surveyed, the death of any seedling should be recorded beside its unique number. New seedlings should be assigned new numbers (do not reassign numbers from dead seedlings) and their height, stem diameter and leaf number recorded.

As seedling density decreases, the size of the study area will have to be increased to retain 25 plants (e.g. by

Figure 6.5 Where to take stem diameter and height measurement

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increasing the width of the belt transect). When the stem diameter is greater than 2.5 cm, measure the stem at a height of 1.3 m, rather than at the base.

6.2.12 Data interpretation The rate of increase in seedling biomass indicates the rate of regeneration of a site. To calculate this, measure the relative volume (as opposed to biomass) of seedlings per square metre within the transect, and its rate of increase over time. Slow or no increase in relative seedling volume and/or high seedling mortality rates may indicate environmental stress.

The relative volume of each seedling within a plot can be calculated from:

Relative volume of seedling (cm3) = x x πD2 x H or x πr2 x H

where:

π = 3.14 (approx.)

D = Diameter of trunk (cm)

H = Height of plant (cm)

r = radius D/2 (cm)

Seedlings differ widely in shape due to their leaves and branches, making true volume difficult to measure. Based on the two key indicators of plant size (stem diameter and height), and the 1/3 multiplication factor, this formula actually measures the volume of the plant stem as if it were pyramid-shaped. This is a relative measurement, allowing the growth rate of seedlings to be monitored and compared with other seedlings.

The total volume of seedlings in the transect is calculated by summing the volumes of all the seedlings measured. Divide the total volume by the area of the transect to give volume (cm3/m2).

Seedling volume cm3/m2 =

where: Σ = Sum of individual seedling volumes

6.2.13 References and further reading Duke, N. (1996) Mangrove reforestation in Panama: an evaluation of planning in areas deforested by a large oil spill, in Field, C. (ed.), Restoration of Mangrove Ecosystems, International Tropical Timber Association (ITTO), Tokyo.

Duke, N. (1992) Aging Rhizophora seedlings from leaf scar nodes: a technique for studying recruitment and growth in mangrove forests, Biotropica, vol. 24, pp. 173-6.

6.3 Canopy cover and leaf area index

6.3.1 Introduction—what is leaf area index and how is it related to mangrove forest health? Leaf area index (LAI) is an index score of the total area of leaf surface within a plant community relative to the ground area of that community. Since plants under stress tend to shed leaves, thus reducing their leaf area, environmental stress can be detected by monitoring LAI. Significant decrease or gradual reduction over time in canopy cover and/or leaf area index score may indicate ecosystem stress or disturbance.

The LAI is not a true measurement of leaf area, but a relative score that can be used to compare results between sites and over time.

LAI can be used to monitor short to long-term foliage patterns and changes in a mangrove stand (e.g. high rates of primary productivity during good seasons, or defoliation through storm damage, seasonal or drought-related leaf fall, insect attack, etc.).

To calculate LAI, the intensity of full sunlight is measured (using a light meter), and this is compared with the light intensity measured under a mangrove canopy. Either a Lux or photosynthetic active radiation (PAR) meter is suitable. The cheap and robust Lux meters measure total light intensity, while the more expensive PAR meters measure photosynthetic active radiation, which is the light absorbed by plants during photosynthesis.

The choice of meter depends on the accuracy required. A scientific approach requires the more expensive and fragile PAR meter, which is highly sensitive and able to detect much smaller foliage pattern changes than the Lux meter can. However, despite some limitations, Lux meters can be used to detect and measure short-term changes

)(m areatransect )(cm volumeseedling relative

2

3Σ

31

41

31

to mangrove canopy cover.

Lux and PAR readings are not easily comparable, so choice of meter is important.

6.3.2 Method summary Light readings are taken in the sun, outside the canopy of a homogenous mangrove community. A series of readings are then taken under the canopy, followed by a further series, again in the sun. LAI is determined by calculating the ratio of light under the canopy, to the light in the adjacent open space.

Best timing is midday ±2 hours to ensure that the sun is as close to overhead as possible, and valid measurements can only be made on sunny days.

LAI measurement is not suitable for fringe mangrove environments, as light penetration from the edges will bias the result (e.g. on narrow river fringes).

6.3.3 Site selection This method involves monitoring the LAI in wide, homogenous stands of mangroves. Prior to site selection, examine aerial photographs and look for patterns of zonation.

Select potential monitoring sites where mangrove forests are homogenous and are wide enough to minimise light infiltration from the forest edges.

At the site, walk through the forest and closely observe its structure (i.e. canopy height, species composition, and stem density) then select an area that appears to be representative of the mangrove community to be monitored. To avoid light penetration, ensure that there are at least 20 m between the quadrat and a forest edge.

Using the 50 m tape, mark out a quadrat of approximately 25 m x 25 m.

6.3.4 Using the light meter Hold the light meter in one hand and the sensor in the other (see Figure 6.6), ensuring that the white surface of the sensor is facing upwards.

Turn on the light meter and select a range that is appropriate for the current light conditions.

To take a reading, blink your eyes and record the first reading that you see when you open them. A second person should carry the datasheets and record the results.

The light meter may have different range settings (e.g. xl, x10 and x100) to allow for a range of light intensities. When recording results, always record the range settings, or make the necessary corrections while taking measurements.

Wipe down the meter with a moist cloth after each use, treating the sensor with extreme care.

Always read instruction manuals for light meters, as some require the application of a correction factor.

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6.3.4.1 Take the light meter readings

Take light readings outside the canopy:

• Turn on the light meter and set the range to 100x. Take five readings outside the canopy (multiplying each by 100 to adjust for range) and record the results.

Take light readings within the quadrat:

• Set the light meter to lx or 10x. Walk along the boundaries of the quadrat taking a light reading every metre for 100 m. Record the results, remembering to adjust for the range.

• When a light meter is used under a forest canopy, readings will occasionally go off-scale. If this happens, switch to a higher range setting and record the measurement on the new scale. Return to the original scale and continue to take readings. It is important to complete each set of readings within 30 minutes.

Take a light reading outside the canopy:

• Set the light meter to 100x and return to the outside of the canopy. Take another five readings and record the results.

6.3.4.2 Measure the zenith angle of the sun

An instrument called a clinometer is used to measure the zenith angle of the sun, which is its angle from the vertical (see Figure 6.7). The closer it is to midday, the smaller this angle will be. If a clinometer is not available, insert a 1-2 m pole into a flat area of sunlit ground, ensuring it is at 90º. Measure the height of the stick and the length of its shadow.

Zenith angle = stick ofHeight

shadow) of(Length Tan -1

Alternatively, the zenith angle for the site can be calculated from a nautical almanac or from a suitable computer program using the latitude, longitude (or GPS reading) and time of day.

Figure 6.6 Using a light meter

Figure 6.7 Zenith angle of the sun

6.3.5 Data interpretation Calculate canopy cover and LAI of a plot using the following formulas:

Canopy cover = readings spaceopen of Average

100 x readingscanopy of Average1 −

LAI = Ln( 0b II ) / -k x cos( 180π∞ )

where:

Ib = Mean value of light below the canopy

I0 = Mean value of light above the canopy

k = is an extinction coefficient that accounts for the angle and orientation of the foliage (A k value of 0.55 has been chosen as appropriate for mangrove stands).

∞ = Zenith angle of the sun

Ln = natural log of number

π = Pi or 3.14

Note: The k value quoted can be used in calculations for closed canopy forests of Rhizophora, Bruguiera and Ceriops spp. but, due to the different structural characteristics of their canopies, is not suitable for use in closed canopy forests of Avicennia spp., or in open forests. However, as no k value has, as yet, been calculated for Avicennia stands, the nominated value can be used to calculate LAI, but the data can be compared only with that from other Avicennia stands.

Forest LAI and canopy cover are the mean results from each plot. Data can be displayed on histograms as the LA1 score, or as canopy cover per plot or forest over time. Median, range and standard deviations of readings are also calculated.

It is important to distinguish between natural and human-induced changes when interpreting data. As leaf area in canopies will naturally vary slightly from season to season, with a peak during the summer months, LAI can also vary naturally between sites and between different communities.

Large reductions in LAI are normally the result of disturbance or stress. If they are detected at a site, compare results from a control or other site (containing the same species) to determine if this reduction is local or more widespread. Volunteers can also return to the site to observe the forest closely for evidence of damage (e.g. storm damage, insect attack or stress).

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6.3.6 References and further reading Clough, B.F., Ong, J.E. & Gong, W.K. (1997) Estimating leaf area index and photosynthetic production in canopies of the mangrove Rhizophora capiculata, Marine Ecology Progress Series, vol. 159, pp. 285–92.

English, S., Wilkinson, C. & Baker, V. (eds) (1997) Survey Manual for Tropical Marine Resources, 2nd edition, Australian Institute of Marine Science, Townsville.

Gordon, D.M., Bougher, A.R., LeProvost, M.I. & Bradley, J.S. (1995) Use of models for detecting and monitoring change in a mangrove ecosystem, North-Western Australia, Environment International, vol. 21, no. 5, pp. 605–18.

6.4 Mangrove forest structure

6.4.1 Introduction—what is mangrove forest structure and how is it related to mangrove forest health? Mangrove structure refers to the composition of a mangrove community in terms of canopy height, stem density, age, tree diameter and species represented. It varies considerably between different forest types, and between the same forest types in different locations. It is influenced by many natural factors including climate, tidal inundation, soil pH and salinity, sediment particle size and amount of freshwater.

Mangrove structure is likely to be affected when any of these parameters is altered (positively or negatively) by human-induced impacts. Positive changes can result in greater forest vigour (increased diameter, canopy cover and stem density), while negative changes can stress the mangrove community, resulting in reduced canopy cover, stem density, tree mortality and, eventually, reduced basal area of trees and/or lower canopy height.

Higher proportions of dead versus live stems, and/or decline in the basal area of the trunks of mangroves in a stand may indicate stress or past disturbance. Many aspects of mangrove structure tend to respond more slowly than most other estuarine indicators, and it can take several years before changes can be detected.

If saltmarshes are present, record the species types and estimate their percentage cover within the quadrat. Inspect the leaves of each tree for any signs of discoloration, wilting or insect damage. If there is insect damage, estimate the amount in terms of percentage loss of leaf surface area for the entire tree. If there are introduced vines, record the species (or describe their features) and estimate their percentage cover.

Also note any evidence of unusual occurrences, such as deposition of rubbish, or other human-induced disturbances.

6.4.2 Method summary This method is used to provide baseline data on the diversity and structure of a mangrove community at a particular site, and to monitor long-term changes and provide a quantitative measure of species composition, stem density, and basal area of trees. This information can be useful for interpreting other parameters, such as leaf trapping ability and LAI. Changes to basal area, stem density and canopy cover can be indicators of ecosystem health.

The method is similar to the mangrove line transects method (English, Wilkinson & Baker 1997), but with some simplifications. Despite these, it is still a difficult, time-consuming method, and should be attempted only to study long-term changes to mangrove forests.

A transect running at right angles from the sea to the land is established, with 10 m x 10 m quadrats in each forest zone along the transect. Within each quadrat, the canopy cover, species type, tree height, sapling/seedling number and stem diameter are recorded.

6.4.3 Site selection Though site selection will depend on the objectives of the monitoring program, in most instances it will be necessary to select sites that are representative of the mangroves in the area. Use aerial photographs to determine the size and extent of the site, and look for zonation patterns between the seaward and landward margins. Sites should be ground-truthed to confirm zonation patterns, and to ensure they are representative.

As mangrove systems are extremely diverse and can vary considerably in structure and floristics over short distances, a person of suitable experience should assist with this process.

6.4.4 Lay out transect and set up quadrats If the program involves monitoring each homogenous zone, establish a transect beginning at right angles to the

seaward side of the mangrove forest, and running to the landward edge (see Figure 6.8). Use the compass to establish the bearing to follow. Identify the major forest types or zones along the transect. For each forest type, find an area to the left of the transect that is representative (in terms of floristics and structure) of that mangrove community. If two quadrats are to be established, ensure that they are at least 20 m apart.

If monitoring a homogenous forest type or a narrow mangrove fringe along a creek, transects can be established parallel to the shoreline. Quadrats can be placed where the forest is representative of the mangrove community, or at regular intervals.

To set up a quadrat, mark out a 10 m x 10 m area within the forest. Use the compass to ensure that the corners are at 90 degrees, and mark each with a PVC pole (see Figure 6.9). Ensure that there is a minimum of 25 trees in each quadrat by increasing or decreasing its size if necessary (e.g. to 5 m x 5 m in a dense forest).

Figure 6.8 How to establish a quadrat and record position of tree using x/y coordinates

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Figure 6.9 How to establish a transect

6.4.5 Estimate canopy cover Imagine that the quadrat has been divided into four smaller quadrats. Stand in the middle of each of these imaginary quadrats and estimate the amount of sky that is blocked by the canopy. This score is referred to as canopy cover. (The illustrations in Figure 6.10 can be used as a guide.)

Calculate the average of these four estimates and record the result.

Note: If a light meter or forest densitometer is available, use this in preference to making visual estimates. See Section 6.3.4 for instructions on measuring canopy cover using a light meter.

Figure 6.10 Estimating canopy cover

6.4.6 Estimate canopy dominance If the canopy consists of more than one species of mangrove, estimate the percentage that each species contributes to the total canopy cover.

Note: Dominance is not the same as canopy cover; the total of all species must equal 100 per cent. For example, if there is a 70 per cent canopy cover and only one species, canopy dominance by that species will be 100 per cent.

6.4.7 Measure stem diameter Measure the stem diameter of each tree at breast height (1.3 m above the ground) (see Figure 6.11). Measure only those trees with a stem diameter of 2.5 cm or more; do not measure saplings or seedlings (see Table 6.2).

Table 6.2 Criteria for distinguishing growth status

Tree Stem DBH (1.3 m) 2.5 cm or greater

Sapling Plant 1 m or more in height, with DBH < 2.5 cm

Seedling Plant < 1 m in height

A diameter tape measures both circumference (girth) and the calculated diameter. Record result as diameter at breast height (DBH) (see Figure 6.11).

A tape measure measures circumference only. Record this as circumference at breast height, and calculate DBH by dividing this result by π (3.14 approx).

If carrying out long-term monitoring, hammer a galvanised nail (half of its length) into stems 10 cm below where measurements have been taken, to provide a reference point for future measurements. Note this on the datasheet.

Figure 6.11 Measurements recorded at breast of height

6.4.7.1 How to measure irregularly shaped trees

Irregularly shaped trees are very common in mangrove forests. If an irregularity occurs at breast height (see Figure 6.12), use the following procedures to measure diameter:

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• For multiple stems, fork below breast height; where stem diameter is 2.5 cm or greater, measure the diameter of each stem at breast height, and record all results in the same box on the datasheet. Do not count each stem as a separate tree.

• For multiple stems, fork at breast height; take the measurement slightly below the swelling caused by the fork. For buttress roots, take the measurement 30 cm above the uppermost prop root or buttress.

• For trunk swellings, take the measurement slightly above or below the swelling.

Some smaller mangrove forests may be naturally stunted or dwarf-like. In such situations these criteria are not suitable for determining growth status.

6.4.8 Count saplings and seedlings Count the number and record species type of seedlings and saplings within the quadrat. If plants are dense, use a smaller quadrat (size will depend on numbers, but 1 m x 1 m is a starting point), ensuring that the area sampled is representative of the larger quadrat. Estimate the number of seedlings/saplings within the 10 m x 10 m quadrat, based on the results of the smaller quadrat sampling

Figure 6.12 Measuring the stem diameter of irregularly shaped tree

6.4.9 Estimate height To measure the height of each tree, stand the height pole up directly below the highest point of the tree (see Figure 6.13). Estimate the height of the tree to the nearest metre, based on the known length of the pole. Record the result.

Note: As this can be very difficult if the forest canopy is higher than 10 m, use of a clinometer is recommended in such situations.

6.4.10 Soils Collect a sample of the substrate from the quadrat and rub it between your fingers. Record the sediment type based on its feel (e.g. sand, mud, sand/mud, mud/sand). Other information such as pH and salinity can be recorded also.

6.4.11 Tag and record position of trees Since branches can die during long-term monitoring of a plot, attach alloy or stainless steel tags to the larger branches or to the main stem. Use nylon cable or stainless steel wire, ensuring that there is enough slack to allow for growth of the trees.

The position of trees should also be recorded using x/y coordinates (see Figure 6.8).

6.4.12 Measurements needed during re-survey As it is likely that a long time may have elapsed before repeat measurements are made, the original corner marker may have disappeared, but plot boundaries can be located using the tags and the x/y coordinates. If new trees have become established, they should be assigned a new number.

6.4.13 Data interpretation Formal measurements provide quantitative data on the structure or level of ecological development of a mangrove community. Data is expressed as:

• stems (living and dead) per hectare

• basal area (square metres per hectare)

• tree height.

Stems per hectare (stems ha-1) is a measure of the density of living mangrove trees. It is calculated using the formula:

Stems ha-1 = )(mplot theof Area

000 10plot x in stems living ofNumber 2

Stems ha-1 should be calculated for each plot, together with the average for all the plots. The number of dead stems per hectare can also be calculated using the above formula, together with the overall ratio of dead to live stems (total dead stems versus total live stems).

Basal area (BA) of a plant refers to the cross-sectional area of its stem at 1.3 m (breast height). The basal area of a stand (stand BA) is the sum of all stem BAs in the quadrat, and is expressed as square metres per hectare (m2/ha).

Figure 6.13 Using a height pole

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BA is a measure of the size or level of ecological development of a mangrove community. Normally, the higher the BA, the greater the biomass and level of development of a mangrove community.

Basal area for an individual plant is calculated using the following formula:

BA (cm2) = πr 2

where:

r = radius of the stem (cm) = 2(cm) DBH

π = 3.14 (approx)

If the plant has multiple stems, the basal area for the plant will be equal to the sum of the basal areas of the individual stems.

To calculate stand BA, use the following formula:

Stand BA (M2/ha) = )(Mplot theof Area

)(CMplot for theBA 2

2∑

where Σ BA = sum of individual BAs. Increases in BA over time may indicate that the community is still growing and developing. Increases in average canopy height will support this theory. A significant decrease in BA may indicate that disturbance has occurred—a theory that would be supported by an increase in the ratio of dead to live stems.

Average or median tree height can also be calculated to provide an indicator of canopy height. However, tree height measurements are better used to track the progress of individual trees, rather than that of the entire forest.

6.4.14 References and further reading Australian Parks and Wildlife Service (1981) The Kakadu National Park Mangrove Forests and Tidal Marshes, Volume 3, Commonwealth of Australia, Canberra.

Clough, B.F. (ed.) (1979) Mangrove ecosystems in Australia—structure, function and management, Proceedings of the Australian National Mangrove Workshop, Australian Institute of Marine Science, Canberra.

Clough, B.F., Dixon, P. & Dalhaus, 0. (1997 Allometric relationships for estimating biomass in multi-stemmed mangrove trees, Australian Journal of Botany, vol. 45, pp. 1023–31.

English, S., Wilkinson, C. & Baker, V. (1997) Survey Manual for Tropical Marine Resources, 2nd edition, Australian Institute of Marine Science, Townsville.

Greening Australia (2001) Tracking Your Community Vegetation Project, Greening Australia, Brisbane.

Lovelock, K. (1997) Field Guide to Mangroves, Australian Institute of Marine Science, Townsville.

Snedaker, S.C. & Snedaker, J.G. (1984) The Mangrove Ecosystem: Research Methods, UNESCO, Paris.

6.5 Crab burrow counts

6.5.1 Introduction—why monitor crab hole density? Estuarine crabs break down much of the leaf and other organic matter produced by mangrove forests. Their burrows also increase the ratio of soil surface area to air, resulting in some aeration and oxidation of the mostly anoxic mangrove soils. This oxidation can be important for the growth of mangrove plants.

Consequently, changes to the crab population can affect the nutrient cycling and oxidation of intertidal soils, which in turn can affect the productivity of mangroves. Decreased crab populations and associated burrow density can lead to decreased nutrient cycling and soil aeration, and reduced production of surrounding plants.

Data on crab burrow density may complement leaf litter trapping or other mangrove monitoring exercises. Crabs can be sensitive to pollution. Their absence from a mangrove forest may indicate that the site is experiencing human-induced stress.

6.5.2 Method summary The number of crab burrows in a survey area is estimated by counting burrows within 50 cm x 50 cm quadrats. Monitor every three months.

6.5.3 Site selection This method is normally used in association with other methods, but if establishing a new site, ensure that it is in a

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homogenous mangrove forest, in an area representative of the surrounding forest.

6.5.4 Establish transects Establish three parallel 10 m transects through the site, 5 m apart. Mark the beginning and end of each with a peg to assist in locating the site again later.

6.5.5 Count the number of crab burrows Starting at 0 m, place a quadrat to the left of the transect and count the number of crab burrows within it. Burrows on the edge of the quadrat should be counted only if the centre of the hole is within the quadrat.

If crab holes are very numerous, use a 25 cm x 25 cm area of the quadrat and multiply the results by four.

Replace the quadrat and count crab holes every 2 m along the length of the transect.

6.5.6 Data interpretation Data is interpreted as crab holes per square metre (holes m-2).

Results may be highly variable between sites, so establish a baseline burrow density for each site. Long-term trends showing a significant decline in burrow numbers may indicate declining crab numbers and/or that the site is experiencing stress. Since crabs can have multiple burrow entrances and some species have been known to share burrows, the relationship is not linear.

Since crab hole abundance does not equate to absolute crab populations, significant changes in burrow counts would need to be recorded to indicate changes in population.

Crab holes can be covered by sediment plugs at low tide.

6.5.7 References and further reading Moritz-Zimmermann, A. & Comley, B. (2000) Overview and methodologies, Mangrove Monitoring Program, Darwin Harbour, Northern Territory, Ch. 1, Department of Lands, Planning and Environment, Northern Territory.

Part G Monitoring seagrass

7.1 Intertidal percentage cover

7.1.1 Introduction Measurement of percentage cover can provide an early warning of seagrass decline and can be used to monitor the long-term health of seagrasses in a local area.

Significant reduction in percentage cover, or failure to recover after a disturbance, may indicate pressure on a seagrass meadow. Although the rate at which seagrass beds recover from disturbances such as floods is unknown, slow or no recovery may indicate other pressures.

7.1.2 What is seagrass percentage cover? Percentage cover is the percentage of a given area covered by seagrass (i.e. percentage cover is 100 per cent if the entire area is covered).

Percentage cover indicates the status and stage of development of a seagrass meadow. For example, if percentage cover is high and varies little over time, this indicates a stable, well-developed community. On the other hand, a reduction in cover, or increasing incidence of algae or epiphytes, may indicate natural or human-induced stress. Changes in percentage cover over time are also a useful indicator of the rate of recovery after disturbance.

7.2 Method for measuring seagrass percentage cover

7.2.1 Method summary Quadrats (50 cm x 50 cm) are placed at regular intervals along three transects in an intertidal seagrass community. See Figure 7.1 for guidance in the use of transects and quadrats.

Figure 7.1 Using a transect

Percentage cover of seagrass in each quadrat can be rapidly assessed using the percentage cover illustration sheet (see Figure 7.2).

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Figure 7.2 Estimating cover

Rapid visual estimates can be subject to individual observer bias; however, working in pairs can help reduce this. Shoot length and species dominance are recorded, and the percentage cover of algae and epiphytes is estimated.

Frequency of measurement should be a minimum of every three months to account for seasonal trends.

Water temperature, salinity and turbidity are also useful parameters to monitor along with seagrass parameters.

7.2.2 Site selection Walk through a seagrass community to ensure that it is relatively homogenous and evenly shaped, with no sandbanks, mud ridges, or changes in the meadow. Select a 50 m x 50 m site that is representative of seagrass in the area concerned, has low variability, and is not difficult to revisit for future monitoring.

Once a site is selected, record its position using GPS and compass bearings from prominent land features to help relocate it again later.

7.2.2.1 Establishing transects

At least three transects are required at each site (see Figure 7.3). The middle transect at each site need to be permanently marked out. To mark where the middle transect begins, drive a plastic star picket into the ground at the landward edge until the top is only 10 cm above the surface, and attach the subsurface buoy. Record the position of the picket using a GPS.

Take a compass bearing from the site to the sea to ensure that the transect is at right angles to the shore. Holding the 50 m tape, follow the compass bearing (90º) for 50 m, and then take a back bearing (270º) to check your position and adjust if necessary. To mark the other end of the transect, drive a picket into the ground at this point.

Run transects 1 and 3 directly parallel to, and to the left and right of, transect 2. To do so, take a bearing at right angles to transect 2 and walk 25 m in that direction. Take a back bearing to check position and adjust if necessary. Set up transects 1 and 3 in the same way as transect 2.

When setting up the transect, hold the tape with the right hand. Always place the quadrats to the right of the transect, and walk on the left to avoid trampling the survey area. Photographic records of sample quadrats can be kept to complement the monitoring program and to assist with quality assurance. Stand above the quadrat (do not

face the sun); place a quadrat photo labeller above the quadrats at 5 m, 25 m and 45 m, showing the site, transect and quadrat numbers; and take a photo. All monitoring should be conducted at low tide. Try to arrive about one hour before this to set up.

7.2.2.2 Estimate percentage cover and species composition

Place the 50 cm x 50 cm quadrat to the right of the transect at 0 m.

Estimate the percentage cover of seagrass in the quadrat using the illustrations in Appendix H as a guide. Record the result. Record the species of seagrasses in the quadrat, and determine the dominance of each (i.e. the percentage of cover that each species represents). These figures must total 100 per cent in all quadrats where seagrass is present.

7.2.2.3 Measure blade length

Ignoring the tallest 20 per cent of leaves, randomly select 3–5 seagrass blades from within the quadrat and measure their length (in cm) from the base to the end. Calculate the average and record the result.

7.2.2.4 Sediment grain size composition

Collect a sample of the substrate from the quadrat and rub it between your fingers. Based on its feel, estimate the grain composition. Record the result in order of dominance (e.g. sand, mud, sand/mud, mud/sand).

7.2.2.5 Estimate epiphyte cover

Measure the percentage cover of epiphytes (microalgae) on the seagrass blades within the quadrat. This figure should be the percentage of seagrass leaf area covered by epiphytes, not quadrat area.

7.2.2.6 Estimate algal cover

Estimate the percentage cover of both filamentous and macroalgal cover in the quadrat, using the procedure in

Figure 7.3 Overview of transect layer

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step 7.2.2.2.

7.2.2.7 Other information

Record the presence of fauna species (including yabbie or worm holes) and any other features of interest, such as dugong trails.

7.2.2.8 Repeat procedure

Repeat steps 2–6 at 5 m intervals along the transect up to, and including, the 50 m mark (i.e. in 11 quadrats). Repeat procedure for each transect.

7.3 Data interpretation Data is presented as:

• mean percentage cover (per site, not per transect)

• mean algal cover

• mean epiphyte cover of seagrass blades

• mean shoot length.

Calculate the range and standard deviation of each parameter. Data is best presented as graphs or histograms which can be used to show trends over time at, and between, sites.

Bias in estimating percentage cover is a potential problem, particularly when values are low. If photographs of the quadrats have been taken, someone with experience can re-evaluate the estimates and a correction factor can be applied. Mean percentage values can be corrected if the level of error is known.

It is important to understand the natural changes that affect seagrass meadows. Seasonally, percentage cover is likely to be higher during the warmer months and lower during winter. To identify these trends it is necessary to collect data for a minimum of two years (but preferably more).

Significant gains or declines in percentage cover of seagrass at a site indicate that the seagrass bed is undergoing change. The rate of change should be compared with that at nearby sites (preferably ones with similar percentage cover) to determine if it is local or more widespread. A decline in cover may be due to environmental stress, while a gain can indicate recovery of a meadow after a flood event.

When comparing data between sites, it is important to realise that different levels of cover don't always indicate that one site is degraded. Some seagrass meadows may naturally have low cover.

High epiphyte and algal cover may indicate that nutrient enrichment is occurring at the site, particularly if there is freshwater or a local source of nutrient nearby. Values for these parameters are likely to be higher during the summer months.

A combination of blade length and percentage cover can indicate the structure of a seagrass meadow. For example, though a quadrat with very few shoots, but long leaf blades, may have the same percentage cover as one with many more shoots but much shorter blades, the structure of the meadow is very different in each.

7.4 References and further reading Campbell, S.J. & McKenzie, L.J. (2001), Seagrass Watch: Community-Based Monitoring of Seagrass Meadows in Hervey Bay and Whitsunday Regions: 1998-2001, Department of Primarv Industries, Cairns.

Kirkman, H. (1997) Seagrasses of Australia, in Estuaries and the Sea, State of the Environment Technical Paper Series, Environment Australia, Canberra.

McKenzie, L.J., Campbell, S.J. & Roder, C.A. (2001) Seagrass Watch: Manual for Mapping and Monitoring Seagrass Resources by Community (Citizen Volunteers), Department of Primary Industries, Cairns.

Short, F.T. & Coles, R.G. (eds) (2001) Global Seagrass Research Methods, Elsevier Science B.V., Amsterdam.

Appendixes

Appendix H1 Checklist of equipment needed for macroinvertebrate field sampling

Tick when collected

250µm macroinvertebrate sampling net Buckets x 4 Sorting tray x 4 Coarse sieve (10 mm) Tweezers and pipettes x 4 Vials, vial labels and plastic bags Waders 70% methylated spirits Squirt bottles

Water quality equipment

Conductivity meter (ensure charged and calibrated) Dissolved oxygen meter (ensure charged and calibrated) pH meter (ensure charged and calibrated) Turbidity meter (ensure charged and calibrated) Water sample bottles (1 L and 250 mL) General equipment Site, habitat, water quality and macroinvertebrate field sheets Tape measure Waterplant and macroinvertebrate field identification books Current meter, staff and prop GPS Maps Sunscreen Car fridge or esky Drinking water Card table and chairs, beach umbrella or tarp and poles Camera and film Pens, pencils, erasers and clipboards Phone/radio First aid kit Shovel, 4WD recovery kit

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Appendix H2 Queensland Site Information Sheet This sheet can be completed at any stage of the program. Some information may be required prior to the trip. Some is collected during the trip and the rest is done afterwards when the site has been confirmed. This sheet only needs to be completed on the first visit to a site. Note: 1:100 000 topographic maps are used for the following information, where they are available. In some areas of Queensland, only 1:250 000 topographic maps are available. Latitude and longitude—Use GPS (global positioning system) and confirm readings on a 1:100 000 topographic map.

Altitude—Use 1:100 000 topographic maps.

Stream order—Hierarchical ordering system based upon the degree of branching (Strahler 1957). Stream orders should be determined using 1:100 000 scale maps. A second-order stream is formed by the joining of two first order-streams; the junction of two second-order streams forms a third order stream, etc. (see Figure H1).

Slope (m/m). = contour distance (m)

distance of stream between contour lines (m)

e.g. 6.5 km between 20 m and 40 m contour lines = 20m/6500m = 0.0031m/m Distance from source (km)—Distance from the site to longest thread of stream source.

AMTD (km) (adjusted middle thread distance)—The distance from the mouth of the stream (i.e. at the ocean or where it joins another stream) to the site.

Reach—An assessment of where in the catchment a site lies with relation to the watershed. Note that this does not necessarily correlate to the altitude.

Catchment area (km2)—The area of land above the site being assessed from which the water drains towards the stream.

Reference or test assessment—Determined using the Reference Condition Selection Criteria (Table 1 and page 3 of Site Information Sheet).

Nearest rainfall station—Within EHP, a departmental database called DRF is used. The same information can be extracted from the Bureau of Meteorology website <www.bom.gov.au>. Recording the station name eliminates searches for the closest station when there is a need to verify/update data.

Nearest weather station—Temperature information can be extracted from the Bureau of Meteorology website (see above). Again, recording the station name eliminates searches for the closest station when there is a need to verify/update data.

Figure H1 Method used by the department to determine stream order (Strahler 1957)

SITE NUMBER ……………………………………………………………………

SITE NAME ………………………………………………………………..……....

LATITUDE ……………………… LONGITUDE ………………………………

GRID REFERENCE ………………………………………………………………

MAP NAME ……………… MAP NUMBER ……… SCALE …………………

ALTITUDE (m) …………………. STREAM ORDER …………………………

SLOPE (m/m) …………… DISTANCE FROM SOURCE (km) ………………

AMTD (km)……………… REACH upland midland lowland

CATCHMENT AREA (km2) ……………………….

REFERENCE or TEST ASSESSMENT (see last page)…………………….…

NEAREST RAINFALL STATION ………………………………………………

NEAREST WEATHER STATION ………………………………………………

ACCESS DETAILS

Directions…………………………………………………………………………….……………... Property Owner ……………………………… Phone No. ..…………………………

Contact ..……………………………………… Phone No. ..………..…………...…..

Access Instructions ..……………………………………………………………………………..…

.…………………………………………………………………………………….…….………………

Notify before each visit? [ ] Yes [ ] No

Permission required? [ ] Yes [ ] No Key required? [ ] Yes [ ] No

Key available from …………………………………………………………………………………………………………..

QUEENSLAND SITE INFORMATION SHEET

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Mud map of access route

Sketch of reach

No. Reference condition selection criteria Level of impact *

1 Influence of intensive agriculture upstream*

Intensive agriculture is that which involves irrigation, widespread soil disturbance, use of agrochemicals and pine plantations. Dry-land grazing does not fall into this category.

2 Influence of major extractive industry (current or historical) upstream*

This includes mines, quarries and sand/gravel extraction.

3 Influence of major urban area upstream

This will be relative to population size, river size and distance between the site and the impact.

4 Influence of significant point-source waste water discharge upstream*

Exceptions can be made for small discharges into large rivers.

5 Influence of dam or major weir*

Sites within the ponded area of impoundments also fail. Sites failing this criterion automatically fail the overall assessment.

6 Influence of alteration to seasonal flow regime

This may be due to abstraction or regulation further upstream than the coverage by Criterion 5. Includes either an increase or decrease in seasonal flow.

7 Influence of alteration to riparian zone

Riparian vegetation should be intact and dominated by native species.

8 Influence of erosion and damage by stock on riparian zone and banks

Stock damage to the stream bed may be included in this category.

9 Influence of major geomorphological change on stream channel

Geomorphological change includes bank slumping, shallowing, braiding and unnatural aggradation or degradation.

10 Influence of alteration to instream conditions and habitats

This may be due to excessive algal and macrophyte growth, by sedimentation and siltation, by reduction in habitat diversity by drowning or drying out of habitats (e.g. riffles) or by direct access of stock into the river.

SITE ASSESSMENT

/50

* Note: the level of impact at a site will generally decrease as the distance from the source of impact increases.

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Each criterion relates to an aspect of human activity that impacts on freshwater ecosystems, where impact is defined as a ‘change from natural condition’. Each criterion is given a score according to the following categories:

1. Very major impact

2. Major impact

3. Moderate impact

4. Minor impact

5. Indiscernible impact.

Sites are assessed using the total score for the 10 criteria. Those sites that have a total greater than 44 are deemed to be reference sites. Sites that are given a score of 1, 2 or 3 for criterion 5 (no dam or major weir upstream) cannot be reference sites.

Appendix H3 Water Quality Sampling Field Sheet This field sheet is intended to record information about the water quality parameters (DO, conductivity, water temperature, etc.) and factors that may have an influence on the water quality parameters (e.g. adjacent land use, bank erosion, etc.). The information on the sheet refers to the entire site (100 m reach) at the time it is sampled.

The front side of this sheet is a generic departmental water quality sheet which enables EHP officers to record information on the water quality sample collection site and field measurements, as well as noting the relevant samples and paperwork for external laboratory analysis. The reverse side of this sheet records observations of factors that may influence the water quality from the entire reach. Information on the types of macrophytes present can also be recorded as well as any notes.

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Appendix H4 Field Sheets Contact the department for a copy of the following templates:

• macroinvertebrate sampling field sheet

• habitat assessment field sheet

• river bioassessment task sheet

• macroinvertebrate identification tally sheet.

Appendix H5 Keys used for identification of Queensland Macroinvertebrate Fauna

Keys used for identification of Queensland macroinvertebrate fauna

Order Author/editor Year Key

General keys Hawking, J.H. 1999 A preliminary guide to keys and zoological information to identify invertebrates from Australian freshwaters

General keys Merritt, R.W. & Cummins, K.W. 1996 An introduction to the aquatic insects of North

America (third edition)

General keys Hawking, J.H. 1995 Monitoring River Health Initiative taxonomic workshop handbook

General keys CSIRO 1991 The insects of Australia (second edition) Volume 1

General keys CSIRO 1991 The insects of Australia (second edition) Volume 2

General keys CSIRO 1999 Interactive guide to Australian aquatic invertebrates, Edition 2 (CD ROM)

General keys Williams, W.D. 1980 Australian freshwater life

Acarina Harvey, M.S. & Growns, J.E. 1998 A guide to the identification of families of Australian

water mites (Arachnida: Acarina)

Coleoptera Glaister, A. 1999 Guide to the identification of Australian Elmidae larvae (Insecta: Coleoptera)

Coleoptera Watts, C. 1998 Preliminary guide to the identification of adult and larval Dytiscidae and adult aquatic Hydrophilidae

Coleoptera Davis, J. 1998 (Insecta: Coleoptera) A guide to the identification of larval Psephenidae water pennies (Insecta: Coleoptera)

Crustacea Sheil, R. 2000 Cladocera (Crustacea)

Crustacea Wilson, G.D.F. 1999 Phreatoicidae (Isopoda, Crustacea)

Crustacea Bradbury, J. 1999 Described Australian Amphipoda

Crustacea Griggs, J.A., Shiel, R.J. & Croome, R.L. 1999

A guide to the identification of Chydorids (Branchiopoda: Anomopoda) from Australian inland waters

Crustacea Horwitz, P., Knott, B. & Williams, W.D. 1995 A preliminary key to the Malacostracan families

(Crustacea) found in Australian inland waters

Crustacea Horwitz, P. 1995 A preliminary key to the species of Decapoda (Crustacea: Malacostraca) found in Australian inland waters

Crustacea Shiel, R.J. 1995 A guide to identification of rotifers, Cladocerans and Copepods from Australian inland waters

Crustacea De Deckker, P. 1995

Notes to help identify Ostracods from Australian inland waters and a guide to Ostracod dissection; Attempt at keying Australian Ostracods for their identification

Diptera Cranston, P. 1997 Identification guide to the Chironomidae of New South Wales

Diptera 1995 Key to aquatic diptera families

Ephemeroptera Suter, P.J. 1999 Illustrated key to the Australian Caenid nymphs (Ephemeroptera: Caenidae)

Ephemeroptera Suter, P.J. 1997 Preliminary guide to the identification of nymphs of Australian Baetid mayflies (Insecta: Ephemeroptera) found in flowing waters

Ephemeroptera Dean, J.C. & Suter, P.J. 1996 Mayfly nymphs of Australia—a guide to genera

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Hemiptera Moller Andersen, N. & Weir, T.A.

1994 Austrobates rivularis, gen. Et sp. Nov., a freshwater relative of Halobates Eschscholtz (Hemiptera: Gerridae), with a new perspective on the evolution of sea skaters


1994 The Girrine water striders of Australia (Hemiptera: Gerridae): Taxonomy, distribution and ecology


1994 The sea skaters, genus Halobates Eschscholtz (Hemiptera: Gerridae), of Australia: Taxonomy, Phylogeny and Zoogeography

Megaloptera Theischinger, G. 2000 Australian alderfly larvae and adults (Insecta: Megaloptera) a preliminary guide to the identification of larvae and survey of adults of Australian alderflies.

Mollusca Ponder, W.F., Clark, S.A. & Dallwitz, M.J. 2000 Freshwater and estuarine molluscs: An interactive,

illustrated key for New South Wales

Mollusca Miller, A.C., Ponder, W.F. & Clark, S.A. 1999

Freshwater snails of the genera Fluvidona and Austropyrgus (Gastropoda, Hydrobiidae) from northern New South Wales and southern Queensland, Australia.

Mollusca Smith, B.J. 1996

New South Wales and southern Queensland, Australia: Identification keys to the families and genera of bivalve and gastropod molluscs found in Australian inland waters

Mollusca Sheldon, F. & Walker, K.F. 1993 Shell variation in Australian Notopala (Gastropoda:

Prosobranchia: Viviparidae)

Mollusca Smith, B.J. 1992 Zoological catalogue of Australia—non-marine Mollusca

Mollusca Walker, J.C. 1988 Classification of Australian buliniform planorbids (Mollusca: Plumonata)

Mollusca Stoddart, J.A. 1985 Analysis of species lineages of some Australian thiarids (Thiaridae, Prosobranchia, Gastropoda) using the evolutionary species concept

Mollusca Kuiper, J.G.J. 1983 The Sphaeriidae of Australia (extract only)

Mollusc 1966 Studies on Ancylidae

Mollusca Stoddart, J.A. Western Australian Viviparids (Prosobranchia: Mullusca)

Mollusca Brown, D.S. Observations on Planorbinae from Australia and New Guinea

Odonata Hawking, J. & Theischinger, G. 1999

Dragonfly larvae (Odonata)—a guide to the identification of larvae of Australian families and identification and ecology of larvae from New South Wales

Odonata Theischinger, G. 2000 Preliminary keys for the identification of larvae of the Australian Gomphides (Odonata)

Plecoptera Yule, C. 1997 Identification guide to the stonefly nymphs of New South Wales and northern Victoria

Plecoptera Tsyrlin, E. 1999 Preliminary key to mature nymphs of Leptoperla stoneflies in Victoria

Trichoptera St.Clair, R.M. 2000 Preliminary keys for the identification of Australian Caddisfly larvae of the family Leptoceridae

Trichoptera St.Clair, R.M. 2000 Preliminary keys for the identification of Australian Caddisfly larvae of the families Odontoceridae, Kokiriidae and Oeconesidae

Trichoptera Dean, J.C. 2000 Preliminary keys for the identification of Australian Caddisfly larvae of the families Antipodoeciidae, Atriplectididae, Limnephilidae and Plectrotarsidae

Trichoptera Dean, J.C. & St Clair, R.M. 1999 Taxonomy of immatures of selected families of

Ephemeroptera and Trichoptera

Trichoptera Dean, J.C. 1999 Preliminary keys for the identification of Australian Trichoptera larvae of the family Hydropsychidae

Trichoptera Cartwright, D.I. 1998

Preliminary guide to the identification of late instar larvae of Australian Polycentropodidae, Glossosomatidae, Dipsuedopsidae and Psychomyiidae (Insecta: Trichoptera)

Trichoptera Jackson, J. 1998 Preliminary guide to the identification of late instar larvae of Australian Calocidae, Helicophidae and Conoesucidae

Trichoptera St Clair, R.M. 1997

Conoesucidae (Insecta: Trichoptera) – Preliminary guide to the identification of late instar larvae of Australian Philorheithridae, Calamoceratidae and Helicopsychidae (Insecta: Trichoptera)

Trichoptera Dean, J.C. 1997 A preliminary guide to the identification of larval Hydroptilidae (Insecta: Trichoptera)

Trichoptera Wells, A. 1997

A key to species of late instar larvae of Australian Trichoptera (families Dipseudopsidae, Glossosomatidae, Polycentropodidae, Psychomyiidae, Ecnomidae, Philopotamidae and Tasmiidae)

Trichoptera Cartwright, D.I. 1997 Identification of late instar larvae of Australian Trichoptera genera

Trichoptera Dean, J.C., St Clair, R.M., Cartwright, D.I. & Wells, A.

1996

Freshwater snails of the genera Fluvidona and Austropyrgus (Gastropoda, Hydrobiidae) from northern New South Wales and southern Queensland, Australia.

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Appendix H6 List of predictor variables used for the Mark I and Mark II models Predictor variables used for the Mark I and Mark II models

Mark I

Spring edge Spring bed Autumn edge Autumn bed

Depth (m)

Distance from source (km)

Latitude1

Longitude1

Water body description2

Water temperature (°C)

Bedrock (%)

Cobble (%)

Gravel (%)

Latitude1

Macrophyte cover3

Minimum velocity (m/s)


Stream slope (m/m)

Alkalinity (mg/L)


Latitude1

Longitude1

Minimum velocity (m/s)

Stream slope (m/m)

Altitude (m) Cobble (%)

Gravel (%)

Maximum velocity (m/s)


Sand (%) Silt (%)

Stream order

Stream slope (m/m)

Stream wetted width (m)


Mark II

Spring edge Spring pool Autumn edge Autumn pool

Alkalinity (mg/L)

Cobble (%)

Number of habitats

Latitude1

Longitude1

Mean annual rainfall (mm)

Slope (m/m)

Soil class4

Stream order


Range in wet season; monthly

rainfall means (mm) (WETR)

Alkalinity (mg/L) Bedrock (%)

Cobble (%)


Latitude1

Longitude1

Mean dry season monthly

rainfall (mm) (MDMR)

Pebble (%)

Ratio of mean wet to mean dry

season monthly rainfall (mm)

(RAWD)

Reach category5

Stream wetted width (m)

Substrate categories6

1 Decimal degrees. 2 (reflow) 1: some part of water body flowing; 2: No flow—surface area <100 m2; 3: No flow—surface area >100 m2. 3 Measured in the habitat: 0: none (0%); 1: little (1-30%); 2: moderate (31-70%); 3: extensive (>70%). 4 Soil class number (1-11)—attained from GIS map overlay. 5 0: lower; 1: middle; 2: upper. 6 Number of substrate categories in reach (bedrock, boulder, cobble, pebble, gravel, and silt/clay—7 categories in total).

Appendix H7 National taxonomic codes for macroinvertebrate families collected in Queensland

National taxonomic codes for macroinvertebrate families collected in Queensland

Order Family National taxon code Order Family National

taxon code Porifera Porifera IA999999 Amphipoda Melitidae OP099999 Hydrozoa Hydridae IB019999 Isopoda Cirolanidae OR129999 Hydrozoa Clavidae IB029999 Isopoda Sphaeromatidae OR139999 Temnocephalidea Temnocephalidea IF499999 Isopoda Janiridae OR189999 Turbellaria Dugesiidae IF619999 Isopoda Oniscidae OR259999 Nemertea Nemertea IH999999 Decapoda Atyidae OT019999 Nematoda Nematoda II999999 Decapoda Palaemonidae OT029999 Nematomorpha Nematomorpha IJ999999 Decapoda Parasticidae OV019999 Tardigrada Tardigrada IR999999 Decapoda Sundathelphusidae OX519999 Rotifera Rotifer JZ999999 Decapoda Grapsidae OX619999 Gastropoda Viviparidae KG019999 Crustacea Crustacea OZ999999 Gastropoda Hydrobiidae KG029999 Collembola Collembola QA999999 Gastropoda Bithyniidae KG039999 Coleoptera Microsporidae QC039999 Gastropoda Thiaridae KG049999 Coleoptera Carabidae QC059999 Gastropoda Lymnaeidae KG059999 Coleoptera Haliplidae QC069999 Gastropoda Ancylidae KG069999 Coleoptera Hygrobiidae QC079999 Gastropoda Planorbidae KG079999 Coleoptera Noteridae QC089999 Gastropoda Physidae KG089999 Coleoptera Dytiscidae QC099999 Gastropoda Neritidae KG109999 Coleoptera Gyrinidae QC109999 Gastropoda Gastropoda KG999999 Coleoptera Hydrophilidae QC119999 Bivalvia Hyriidae KP019999 Coleoptera Hydraenidae QC139999 Bivalvia Corbiculidae KP029999 Coleoptera Staphylinidae QC189999 Bivalvia Sphaeriidae KP039999 Coleoptera Scirtidae QC209999 Bivalvia Bivalvia KP999999 Coleoptera Elmidae QC349999 Hirudinea Glossiphoniidae LH019999 Coleoptera Limnichidae QC359999 Hirudinea Ozobranchidae LH029999 Coleoptera Heteroceridae QC369999 Hirudinea Richardsonianidae LH039999 Coleoptera Psephenidae QC379999 Hirudinea Ornithobdellidae LH049999 Coleoptera Ptilodactylidae QC399999 Hirudinea Erpobdellidae LH059999 Coleoptera Chrysomelidae QCAH9999 Hirudinea Hirudinea LH999999 Coleoptera Brentidae QCAM9999 Oligochaeta Oligochaeta LO999999 Coleoptera Curculionidae QCAN9999 Polychaeta Polychaeta LP999999 Coleoptera Coleoptera QCZZ9999 Arachnida Acarina MM999999 Diptera Tipulidae QD019999 Anostraca Anostraca OD999999 Diptera Tanyderidae QD039999 Conchostraca Conchostraca OF999999 Diptera Blephariceridae QD049999 Cladocera Cladocera OG999999 Diptera Chaoboridae QD059999 Ostracoda Ostracoda OH999999 Diptera Dixidae QD069999 Copepoda Copepoda OJ999999 Diptera Culicidae QD079999 Branchiura Branchiura OK999999 Diptera Ceratopogonidae QD099999 Amphipoda Talitridae OP019999 Diptera Simuliidae QD109999 Amphipoda Ceinidae OP029999 Diptera Thaumaleidae QD119999 Amphipoda Eusiridae OP039999 Diptera Psychodidae QD129999 Amphipoda Corophiidae OP059999 Diptera Athericidae QD229999 Amphipoda Paramelitidae OP069999 Diptera Tabanidae QD239999 Diptera Stratiomyidae QD249999 Neuroptera Osmylidae QN039999 Diptera Empididae QD359999 Neuroptera Neurorthidae QN049999 Diptera Dolichopodidae QD369999 Neuroptera Sisyridae QN059999 Diptera Syrphidae QD439999 Zygoptera Coenagrionidae QO029999 Diptera Sciomyzidae QD459999 Zygoptera Isostictidae QO039999

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Order Family National taxon code Order Family National

taxon code Diptera Ephydridae QD789999 Zygoptera Protoneuridae QO049999 Diptera Muscidae QD899999 Zygoptera Lestidae QO059999 Diptera s-f Aphroteniinae QDAA9999 Zygoptera Hypolestidae QO069999 Diptera s-f Diamesinae QDAB9999 Zygoptera Megapodagrionidae QO079999 Diptera s-f Telmatogetoninae QDAC9999 Zygoptera Synlestidae QO089999 Diptera s-f Podonominae QDAD9999 Zygoptera Diphlebidae QO099999 Diptera s-f Tanypodinae QDAE9999 Anisoptera Aeshnidae QO129999 Diptera s-f Orthocladiinae QDAF9999 Anisoptera Gomphidae QO139999 Diptera s-f Chironominae QDAJ9999 Anisoptera Petaluridae QO159999 Diptera Chironomidae (unid.) QDAZ9999 Anisoptera Corduliidae QO169999 Diptera Diptera QDZZ9999 Anisoptera Libellulidae QO179999 Ephemeroptera Baetidae QE029999 Zygoptera Zygoptera QO999997 Ephemeroptera Ameletopsidae QE049999 Anisoptera Anisoptera QO999998 Ephemeroptera Leptophlebiidae QE069999 Plecoptera Eustheniidae QP019999 Ephemeroptera Ephemerellidae QE079999 Plecoptera Gripopterygidae QP039999 Ephemeroptera Caenidae QE089999 Plecoptera Plecoptera QP999999 Ephemeroptera Prosopistomatidae QE099999 Trichoptera Hydrobiosidae QT019999 Ephemeroptera Ephemeroptera QE999999 Trichoptera Glossosomatidae QT029999 Hymenoptera Hymenoptera QG999999 Trichoptera Hydroptilidae QT039999 Hemiptera Mesoveliidae QH529999 Trichoptera Philopotamidae QT049999 Hemiptera Hebridae QH539999 Trichoptera Hydropsychidae QT069999 Hemiptera Hydrometridae QH549999 Trichoptera Polycentropodidae QT079999 Hemiptera Veliidae QH569999 Trichoptera Ecnomidae QT089999 Hemiptera Gerridae QH579999 Trichoptera Psychomyiidae QT099999 Hemiptera Saldidae QH609999 Trichoptera Tasimiidae QT139999 Hemiptera Nepidae QH619999 Trichoptera Conoesucidae QT159999 Hemiptera Belostomatidae QH629999 Trichoptera Antipodoeciidae QT169999 Hemiptera Ochteridae QH639999 Trichoptera Helicopsychidae QT179999 Hemiptera Gelastocoridae QH649999 Trichoptera Calocidae/Helicophidae QT189999 Hemiptera Corixidae QH659999 Trichoptera Philorheithridae QT219999 Hemiptera Naucoridae QH669999 Trichoptera Odontoceridae QT229999 Hemiptera Notonectidae QH679999 Trichoptera Atriplectidae QT239999 Hemiptera Pleidae QH689999 Trichoptera Calamoceratidae QT249999 Hemiptera Hemiptera QHZZ9999 Trichoptera Leptoceridae QT259999 Mecoptera Nannochoristidae QK019999 Trichoptera Dipseudopsidae QT269999 Lepidoptera Pyralidae QL019999 Trichoptera Trichoptera QT999999 Lepidoptera Lepidoptera QL999999 Unidentified Unidentified XX999999 Megaloptera Corydalidae QM019999 Megaloptera Sialidae QM029999

Appendix H8 Transport of live aquatic animals

Unaccompanied road transport

Place finfish/crustacea in two strong large plastic bags, with the inner one 1/4 to 1/3 full of water from the pond or tank from which the animals came (Figure H2a). Fill the inner bag with air (for a 1-2 hour trip) or oxygen (for a longer trip). Be sure to expel all air from the inner bag prior to filling with oxygen. Bring the top together, fold back on itself and seal with a rubber band (Figure H2b). Seal the second plastic bag over the top of the inner one (Figure H2c).

Place the bag in a large insulated container, e.g. a polystyrene esky. Label correctly and include written information on the history/problem of the animals it contains (attach to outside).

Accompanied road transport

The above method may be used. Alternatively, a plastic rubbish bin lined with a large unsealed plastic bag may be used. The transported water can be aerated during the trip with a portable battery-operated aquarium air pump or a fisherman's bait aerator.

Air transport

Transport of live animals within a state by air must conform to current Australian Domestic Airline Seafood Regulations and Packaging Approvals and IATA regulations as specified by the airline. The regulations outline the correct packing procedures and the approved transport box to be used. For example, live freshwater fish must be packed and transported according to IATA regulations, while the transport of live saltwater fish must comply with the current Australian Domestic Airline Seafood Regulations and Packaging Approvals. Contact the freight section of your nearest airport to check requirements before packing and transporting live fish to the laboratory.

NOTE: Interstate transport of sick animals is illegal, since there may be a risk of introducing a disease to a state declared free of a particular disease. You must contact your nearest Animal Health Laboratory within Queensland if you have a suspected disease outbreak.

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Glossary

Abiotic The non-living components of a system (see biota).

Absorption In chemistry: penetration of one substance into the body of another. In biology: the act of absorbing (i.e. to take in as fluids or gases through a cell membrane). To take a substance (e.g. water, nutrients) into the body through the skin or mucous membranes or, in plants, through root hairs. See also Adsorption.

Acidic Having a high hydrogen ion concentration (low pH).

Acid-soluble metal The concentration of a metal that passes through a 0.45 micron (μm) membrane filter after the sample is acidified to pH 1.5–2.0 with nitric acid.

Acid volatile sulphide (AVS) Sulfides in sediment that liberate hydrogen sulphide on reaction with cold dilute acid (mainly FeS or MnS in sediments).

Acute toxicity Rapid adverse effect (e.g. death) caused by a substance in a living organism. Can be used to define either the exposure or the response to an exposure (effect).

Adsorption The taking up of one substance at the surface of another.

Aerobic Organisms requiring oxygen for respiration or conditions where oxygen is available.

Algae Comparatively simple chlorophyll-bearing plants, most of which are aquatic and microscopic in size.

Alkalinity The quantitative capacity of aqueous media to react with hydroxyl ions. The equivalent sum of the bases that are titratable with strong acid. Alkalinity is a capacity factor that represents the acid neutralising capacity of an aqueous system.

Ambient waters All surrounding waters, generally of largely natural occurrence.

Anaerobic Conditions where oxygen is lacking (anoxic); organisms not requiring oxygen for respiration.

Analyst A particular person in the receiving laboratory who is to perform or supervise the analyses on the samples in question and has recognised qualifications.

Analytes The physical and chemical species (indicators) to be determined.

Antagonism A phenomenon in which the effect or toxicity of a mixture of chemicals is less than that which would be expected from a simple summation of the effects or toxicities of the individual chemicals present in the mixture.

Anthropogenic Produced or caused by humans.

Aquatic ecosystem Any watery environment from small to large, from pond to ocean, in which plants and animals interact with the chemical and physical features of the environment.

Aquifer An underground layer of permeable rock, sand or gravel that absorbs water and allows it free passage through pore spaces.

Assimilation The incorporation of absorbed substances into cellular material.

Assimilative capacity The maximum loading rate of a particular pollutant that can be tolerated or processed by the receiving environment without causing significant degradation to the quality of the ecosystem and hence the environmental values it supports.

Baseline data Also called pre-operational data (studies); collected (undertaken) before a development begins.

Benthic Referring to organisms living in or on the sediments of aquatic habitats (lakes, rivers, ponds, etc.).

Benthos The sum total of organisms living in, or on, the sediments of aquatic habitats.

Bioaccumulation General term describing a process by which chemical substances are accumulated by aquatic organisms from water, either directly or through consumption of food containing the chemicals.

Bioassay A test that exposes living organisms to several levels of a substance that is under investigation, and evaluates the organism’s responses.

Bioavailable The fraction of the total of a chemical in the surrounding environment that can be taken up by organisms. The environment may include water, sediment, soil, suspended particles, and food items.

Biochemical (or Biological) Oxygen Demand (BOD)

The decrease in oxygen content in mg/L of a sample of water in the dark at a certain temperature over a certain of period which is brought about by the bacterial breakdown of organic matter. Usually the decomposition has proceeded so far after 20 days that no further change occurs. The oxygen demand is measured after five days (BOD5), at which time 70% of the final value has usually been reached.

Bioconcentration factor (BCF) A unitless value describing the multiple by which a chemical can be concentrated in the tissues of an organism in the aquatic environment. At apparent equilibrium during the uptake phase of a bioconcentration test, the BCF is the concentration of a chemical in one or more tissues of the aquatic organisms divided by the average exposure concentration in the test.

Biodiversity (biological diversity) The variety of life forms, including the plants, animals and microorganisms, the genes they contain and the ecosystems and ecological processes of which they are a part.

Biofilm Layer of materials created by microorganisms on an underwater surface.

Biological assessment Use and measurement of the biota to monitor and assess the ecological health of an ecosystem.

Biological community An assemblage of organisms characterised by a distinctive combination of species occupying a common environment and interacting with one another.

Biomagnification Result of the processes of bioconcentration and bioaccumulation by which tissue concentrations of bioaccumulated chemicals increase as the chemical passes up through two or more trophic levels. The term implies an efficient transfer of chemicals from food to consumer, so that residue concentrations increase systematically from one trophic level to the next.

Biomass The living weight of a plant or animal population, usually expressed on a unit area basis.

Biosolids Sewage sludge, organic residuals remaining after domestic sewage treatment.

Biota The sum total of the living organisms of any designated area.

Bioturbation The physical disturbance of sediments by burrowing and other activities of organisms.

Bivalve A mollusc with a hinged double shell.

Bloom An unusually large number of organisms per unit of water, usually algae, made up of one or a few species.

Buffer A solution containing a weak acid and its conjugate weak base, the pH of which changes only slightly on the addition of acid or alkali.

Buffering capacity A measure of the relative sensitivity of a solution to pH changes on addition of acids or base.

°C Degrees Celsius.

Carcinogen A substance that induces cancer in a living organism.

Catchment The total area draining into a river, reservoir, or other body of water.

Chemical oxygen demand (COD) The amount of oxygen required to oxidise all organic matter that is susceptible to oxidation by a strong chemical oxidant.

Chlorination • The process of introducing one or more chlorine atoms into a compound. • The application of chlorine to water, sewage or industrial wastes for disinfection.

Chronic Lingering or continuing for a long time; often for periods from several weeks to years. Can be used to define either the exposure of an aquatic species or its response to an exposure (effect). Chronic exposure typically includes a biological response of relatively slow progress and long continuance, often affecting a life stage.

Chronic value The geometric mean of the lower and upper limits obtained from an acceptable chronic test or by analysing chronic data using a regression analysis. A lower chronic limit is the highest tested concentration that did not cause an unacceptable amount of adverse effect on any of the specified biological measurements, and below which no tested concentration caused unacceptable effect. An upper chronic limit is the lowest tested concentration that did cause an unacceptable amount of adverse effect on one or more biological measurements and above which all tested concentrations also caused such an effect.

Colloid Material in solution typically 1 nm–100 nm in diameter. Colloidal particles do not settle out of solution through the force of gravity. Organic colloidal matter is considered especially important in the transport of inorganic substances such as P through the soil profile.

Community An assemblage of organisms characterised by a distinctive combination of species occupying a common environment and interacting with one another.

Community composition All the types of taxa present in a community.

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Community metabolism The biological movement of carbon in an ecosystem, involving two processes, production (via photosynthesis) and respiration.

Community structure All the types of taxa present in a community and their relative abundances.

Compliance Action in accordance with upholding a ‘standard’ (water quality).

Concentration The quantifiable amount of chemical in, say, water, food or sediment.

Criteria (water quality) Scientific data evaluated to derive the recommended quality of water for different uses.

Cyanobacteria A division of photosynthetic bacteria, formerly known as blue-green algae that can produce strong toxins.

Cytotoxic Having an adverse impact on living cells.

Decision criteria Criteria by which decisions will be made as a result of monitoring for potential impacts.

Depuration Process whereby the re is elimination of contaminant from the tissues when an organism is surrounded by uncontaminated water or sediment; also used to describe the use of a controlled aquatic environment to reduce the level of pathogenic organisms that may be present in live shellfish prior to marketing.

Detection limit The smallest concentration or amount of a substance that can be reported as present with a specified degree of certainty by definite complete analytical procedures.

Detritus Unconsolidated sediments composed of both inorganic and dead and decaying organic material.

Dinoflagellates Major class of marine algae that move by flagella. They are often red in colour, and can produce strong toxins that can kill many fish and other marine organisms.

Direct toxicity assessment (DTA) A laboratory procedure for quantifying the potential toxicity of a sample of effluent through exposing a range of test specimens to that effluent. It assesses the toxicity of mixtures of bioavailable chemicals rather than individual chemicals. Also known as Whole Effluent Toxicity (WET) testing.

Diurnal Daily.

Dose The quantifiable amount of a material introduced into an animal.

EC50 (or median effective concentration) The concentration of material in water that is estimated to produce a measurable response in 50% of the test organisms. The EC50 is usually expressed as a time-dependent exposure value (e.g. 24-hour or 96-hour EC50). See also LC50.

Effect size The size of impact that would cause concern (or constitute an early warning). Often defined as a level of (ecological) change that is acceptable in comparison to a defined reference.

Effluent A complex waste material (e.g. liquid industrial discharge or sewage) discharged into the environment.

Electrical conductivity The ability of water or soil solution to conduct an electric current.

End-points Measured attainment response, typically applied to ecotoxicity or management goals.

Endemic, endemism Confined in occurrence to a local region.

Enterococci Any streptococcal bacteria normally found in the human intestinal tract; usually non-pathogenic.

Environmental authority or development approval

A licence, permit or other authority for an environmentally relevant activity, granted by an administering body in accordance with the Environmental Protection Act 1994.

Environmental values Particular values or uses of the environment that are important for a healthy ecosystem or for public benefit, welfare, safety or health and that require protection from the effects of pollution, waste discharges and deposits. Several environmental values may be designated for a specific water body.

Ephemeral stream A stream that carries water only during or immediately after periods of rainfall.

Epilimnion The uppermost layer of water in a lake, characterised by an essentially uniform temperature that is generally warmer than elsewhere in the lake, and by relatively uniform mixing by wind and wave action.

Epilithon Organisms attached to rocks, such as algae and lichens.

Epiphyte A plant that grows on the outside of another plant, using it for support only and not obtaining food from it.

Eukaryotes An organism characterised by the presence of membrane-bound organelles. See also Prokaryote.

Euphotic Of surface waters to a depth of approximately 80–100 m; the lit region that extends virtually from the water surface to the level at which photosynthesis fails to occur because of reduced light penetration.

Euryhaline Describes organisms that are capable of osmoregulating over a wide range of salinities.

Eutrophic Abundant in nutrients and having high rates of productivity frequently resulting in oxygen depletion below the surface layer of a water body.

Eutrophication Enrichment of waters with nutrients, primarily phosphorus, causing abundant aquatic plant growth and often leading to seasonal and/or diurnal deficiencies in dissolved oxygen.

Fate Disposition of a material in various environmental compartments (e.g. soil or sediment, water, air, biota) as a result of transport, transformation and degradation.

Flocculation • The process by which suspended colloidal or very fine particles coalesce and agglomerate into well-defined hydrated floccules of sufficient size to settle rapidly.

• The stirring of water after coagulant chemicals have been added to promote the formation of particles that will settle.

Flow-through system An exposure system for aquatic toxicity tests in which the test material solutions and control water flow into and out of test chambers on a once-through basis either intermittently or continuously.

Fouling Accumulation of material through chemical, physical or biological processes.

Gastropod A mollusc of the class Gastropoda, with a locomotive organ placed ventrally (e.g. snail and limpet).

Gross alpha (activity) A measure of the concentration of alpha-particle emitting radionuclides in water. This is determined by standard techniques involving the evaporation of a water sample and measurement of the alpha activity of the residue.

Gross beta (activity) A measure of the concentration of beta-particle emitting radionuclides in water. This is determined by standard techniques involving the evaporation of a water sample and measurement of the beta activity of the residue.

Groundwater Water stored underground in rock crevices and in the pores of geologic materials that make up the earth's crust; water that supplies springs and wells. See also Aquifer.

Guideline trigger values These are the concentrations (or loads) of the key performance indicators measured for the ecosystem, below which there exists a low risk that adverse biological (ecological) effects will occur. They indicate a risk of impact if exceeded and should ‘trigger’ some action, either further ecosystem specific investigations or implementation of management/remedial actions.

Guideline (water quality) Numerical concentration limit or narrative statement recommended to support and maintain a designated water use.

Habitat The place where a population (e.g. human, animal, plant, microorganism) lives and its surroundings, both living and non-living.

Half-life Time required to reduce by one-half the concentration of a material in a medium (e.g. soil or water) or organism (e.g. fish tissue) by transport, degradation, transformation or depuration.

Hardness The concentration of all metallic cations, except those of the alkali metals (e.g. sodium and potassium), present in water. In general, hardness is a measure of the concentration of calcium and magnesium ions in water and is frequently expressed as mg/L calcium carbonate equivalent.

Hazard The potential or capacity of a known or potential environmental contaminant to cause adverse ecological effects.

High reliability guideline trigger values

Trigger values that have a higher degree of confidence because they are derived from an adequate set of chronic toxicity data and hence require less extrapolation from the data to protect ecosystems.

Humic substances Organic substances only partially broken down that occur in water mainly in a colloidal state. Humic acids are large-molecule organic acids that dissolve in water.

Hydrolysis • The formation of an acid and a base from a salt by the ionic dissociation of water. • The decomposition of organic compounds by interaction with water.

Hydrophilic Having an affinity for water, readily absorbs water.

Hydrophobic Having little or no affinity for water, repels or does not absorb water.

Hypolimnion The region of a water body that extends from below the thermocline to the bottom of the lake; it is thus removed from much of the surface influence.

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Hypothesis Supposition made from known facts as a starting point for further investigation.

Hypoxia Deficiency of oxygen in tissues or in blood; anoxia.

Indicator A parameter that can be used to provide a measure of the quality of water or the condition of an ecosystem.

Inorganic carbon Generally, simple ions and molecules that contain carbon bonded only to inorganic atoms. Carbonates are the most common group, although the cyanide ion is also considered to be inorganic.

Intermittent stream A stream that carries water a considerable portion of the time, but that ceases to flow occasionally or seasonally because bed seepage and evapotranspiration exceed the available water supply.

Interstitial Occurring in interstices or spaces; applied to water and to flora and fauna living between sand grains and soil particles.

Invertebrates Animals lacking a dorsal column of vertebrae or a notochord.

In vitro Outside the intact organism; generally applied to experiments involving biochemical events occurring in tissue fragments or fractions in a laboratory.

Ion An electrically charged atom.

LC100 Lowest concentration of a toxicant that kills all the test organisms.

LC50 (or median lethal concentration) The concentration of material in water that is estimated to be lethal to 50% of the test organisms. The LC50 is usually expressed as a time-dependent exposure value, e.g. 24-hour or 96-hour LC50, the concentration estimated to be lethal to 50% of the test organisms after 24 or 96 hours of exposure.

LD50 (or median lethal dose) The dose of material that is estimated to be lethal to 50% of the test organisms. Appropriate for use with test animals such as rats, mice and dogs, it is rarely applicable to aquatic organisms because it indicates the quantity of a material introduced directly into the body by injection or ingestion rather than the concentration of the material in water in which aquatic organisms are exposed during toxicity tests.

Leachate Water that has passed through a soil and that contains soluble material removed from that soil.

Lethal Causing death by direct action. Death of aquatic organisms is the cessation of all visible signs of biological activity.

Level of protection A level of quality desired by stakeholders and implied by the selected management goals and water quality objectives for the water resource.

Life-cycle study A chronic (or full chronic) study in which all the significant life stages of an organism are exposed to a test material. Generally, a life-cycle test involves an entire reproductive cycle of the organism.

Live weight Weight of the living animal.

LOEC (or lowest observed effect concentration) The lowest concentration of a material used in a toxicity test that has a statistically significant adverse effect on the exposed population of test organisms as compared with the controls.

LOEL (or lowest observed effect level) The lowest concentration that produces an observable effect in a test species. Below this concentration there are no observed effects in the test species.

Long-term trigger value (LTV) The maximum concentration of contaminant in irrigation water which can be tolerated assuming 100 years of irrigation, based on key irrigation loading assumptions.

Low reliability guideline trigger values

Trigger values that have a low degree of confidence because they are derived from an incomplete data set. They are derived using either assessment factors or from modelled data using the statistical method. They should only be used as interim indicative working levels.

Macrophyte A member of the macroscopic plant life of an area, especially of a body of water; large aquatic plant.

Median Middle value in a sequence of numbers.

Mesotrophic Water bodies or organisms which are intermediate between nutrient-rich and nutrient-poor.

Metabolite Any product of metabolism.

Mixing zones An explicitly defined area around an effluent discharge where effluent concentrations may exceed guideline values and therefore result in certain environmental values not being protected. The size of the mixing zone is site-specific.

Moderate reliability guideline trigger values

Trigger values that have a moderate degree of confidence because they are derived from an adequate set of acute toxicity data and hence require more extrapolation than high reliability trigger values, including an acute-to-chronic conversion.

Neurotoxin Toxic substances which adversely affect the nervous system.

NOEC (or no observed effect concentration) The highest concentration of a toxicant at which no statistically significant effect is observable, compared to the controls; the statistical significance is measured at the 95% confidence level. Not detectable or below the limit of detection of a specified method of analysis.

Octanol: water partition coefficient (Kow)

The ratio of a chemical's solubilities in n-octanol and water at equilibrium. Kow is used as an indication of a chemical's propensity for bioconcentration by aquatic organisms where n-octanol is a surrogate for typical animal lipid. The symbol Pow is sometimes used in place of Kow but they are alternative names for the same equilibrium solubility ratio.

Oligotrophic Waters with a small supply of nutrients.

Organic carbon Generally carbon which is chemically bonded to other carbon atoms, although methane (one carbon atom only) and its derivatives are considered organic.

Organism Any living animal or plant; anything capable of carrying on life processes.

Osmoregulation The biological process of maintaining the proper salt concentration in body tissues to support life.

Osmosis Diffusion of a solvent through a semi-permeable membrane into a more concentrated solution, tending to equalise the concentrations on both sides of the membrane.

Oxidation The combination of oxygen with a substance, or the removal of hydrogen from it or, more generally, any reaction in which an atom loses electrons.

Oxygenation The process of adding dissolved oxygen to a solution.

PAH Polycyclic aromatic hydrocarbons.

Parameter A measurable or quantifiable characteristic or feature.

Partition coefficient A ratio of the equilibrium concentration of the chemical between media, for example, between a non-polar and polar solvent, or between air and water, or water and sediment.

Pathogen An organism capable of eliciting disease symptoms in another organism.

Pelagic Term applied to organisms which inhabit the open seas and oceans.

Percentile Division of a frequency distribution into one hundredths.

Periphyton The organisms attached to submerged plants.

Pesticide A substance or mixture of substances used to kill unwanted species of plants or animals.

pH Value that represents the acidity or alkalinity of an aqueous solution. It is defined as the negative logarithm of the hydrogen ion concentration of the solution.

Photodegradation Breakdown of a substance by exposure to light; the process whereby ultra-violet radiation in sunlight attacks a chemical bond or link in a chemical structure.

Photolysis The decomposition of a compound into simpler units as a result of the absorption of one or more quanta of radiation.

Photosynthesis The conversion of carbon dioxide to carbohydrates in the presence of chlorophyll using light energy.

Physico-chemical Refers to the physical (e.g. temperature, electrical conductivity) and chemical (e.g. concentrations of nitrate, mercury) characteristics of water.

Phytoplankton Small (often microscopic) aquatic plants suspended in water.

Phytotoxicity Toxicity of contaminants to plants.

Plankton Plants (phytoplankton) and animals (zooplankton), usually microscopic, floating in aquatic systems.

Pollution The introduction of unwanted components into waters, air or soil, usually as a result of human activity, e.g. hot water in rivers, sewage in the sea, oil on land.

Polychlorinated biphenyls (PCBs) Highly toxic and persistent compounds derived from the replacement of numerous H radicals by Cl radicals on biphenyl, which consists of two benzene rings joined by a covalent bond, with the elimination of two H radicals (C12H10).

Potable water Water suitable, on the basis of both health and aesthetic considerations, for drinking or culinary purposes.

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Practical quantitation limit (PQL) The best level achievable among laboratories within specified limits during routine laboratory operations. The PQL represents a practical and routinely achievable detection level with a relatively good certainty that any reported value is reliable. The PQL is often around five times the method detection limit.

Precipitation • The formation of solid particles in a solution; generally, the settling out of small particles.

• The settling-out of water from cloud, in the form of rain, hail, snow, etc.

Primary production The production of organic matter from inorganic materials.

Producers Organisms that are able to build up their body substance from inorganic materials.

Prokaryotes Organisms characterised by the absence of membrane-bound organelles (opposite to eukaryotes).

Protocol A formally agreed method and procedure for measuring an indicator; it defines the sampling, sample handling and sample analysis procedures.

Protozoans Single-celled, animal-like organisms of the kingdom Protista.

Quality assurance (QA) The implementation of checks on the success of quality control (e.g. replicate samples, analysis of samples of known concentration).

Quality control (QC) The implementation of procedures to maximise the integrity of monitoring data (e.g. cleaning procedures, contamination avoidance, sample preservation methods).

Redox potential An expression of the oxidising or reducing power of a solution relative to a reference potential. Redox potential is dependent on the nature of the substances dissolved in the water, as well as on the proportion of their oxidised and reduced components.

Reference condition An environmental quality or condition that is defined from as many similar systems as possible and used as a benchmark for determining the environmental quality or condition to be achieved and/or maintained in a particular system of equivalent type.

Release of water Water freed from a site for specified purposes such as disposal. May occur by means of infrastructure such as a constructed channel or pipe, or by a natural flow pathway such as an overland gully or drainline.

Residence time The period of time that a volume of liquid (and any associated contaminants) remains in a waterway, catchment system, creek bed, or a part thereof.

Salinity The presence of soluble salts in or on soils or in water.

Sediment Unconsolidated mineral and organic particulate material that settles to the bottom of aquatic environment.

Sediment pore waters Water that occupies the space between particles in a sediment, as distinct from overlying water which is the water above the sediment layer.

Sewage fungus A thick filamentous bacterial growth that develops in water contaminated with sewage. The filamentous material is composed predominately of the bacterium Sphaerotilus natans.

Short-term trigger value (LTV) The maximum concentration of contaminant in irrigation water which can be tolerated for a shorter period of time (20 years) assuming the same maximum annual irrigation loading to soil as for the long-term trigger value.

Sodicity The presence of a high proportion of sodium ions relative to other cations in a soil.

Sorption Process whereby contaminants in soils adhere to the inorganic and organic soil particles.

Speciation The distribution of an element among defined chemical forms, for example, valency or organic and inorganic forms.

Species A group of organisms that resemble each other to a greater degree than members of other groups and that form a reproductively isolated group that will not produce viable offspring if bred with members of another group.

Species richness The number of species present (generally applied to a sample or community).

Suspended particulate matter (SPM)

This is insoluble material which resides in the water column, or is dispersed in a sample upon agitation.

Standard (water quality) An objective that is recognised in enforceable environmental control laws.

Sub-lethal Involving a stimulus below the level that causes death.

Supersaturation Refers to a solution containing more solute than equilibrium conditions will allow.

Synergism A phenomenon in which the effect or toxicity of a mixture of chemicals is greater than that to be expected from a simple summation of the effects or toxicities of the individual chemicals summation of the effects or toxicities of the individual chemicals present in the mixture.

Teratogen An agent that increases the incidence of congenital malformations.

Thermotolerant coliforms Also known as faecal coliforms. In tropical and sub-tropical areas, thermotolerant coliforms may on some occasions include microorganisms of environmental rather than faecal origin.

Threshold concentration A concentration above which some effect (or response) will be produced and below which it will not.

Total dissolved solids (TDS) A measure of the inorganic salts (and organic compounds) dissolved in water.

Total metal The concentration of a metal in an unfiltered sample that is digested in strong nitric acid.

Toxicant A chemical capable of producing an adverse response (effect) in a biological system at concentrations that might be encountered in the environment, seriously injuring structure or function or producing death. Examples include biocides, pesticides and biotoxins (i.e. domoic acid, ciguatoxin and saxitoxins).

Toxicity The inherent potential or capacity of a material to cause adverse effects in a living organism.

Toxicity identification and evaluation (TIE)

Toxicity characterisation procedures involving use of selective chemical manipulations or separations and analyses coupled with toxicity testing to identify specific classes of chemicals and ultimately individual chemicals that are responsible for the toxicity observed in a particular sample.

Toxicity test The means by which the toxicity of a chemical or other test material is determined. A toxicity test is used to measure the degree of response produced by exposure to a specific level of stimulus (or concentration of chemical).

Trigger values These are the concentrations (or loads) of the key performance indicators measured for the ecosystem, below which there exists a low risk that adverse biological (ecological) effects will occur. They indicate a risk of impact if exceeded and should ‘trigger’ some action, either further ecosystem-specific investigations or implementation of management/remedial actions.

Volatile Having a low boiling or subliming pressure (a high vapour pressure).

Water quality criteria Scientific data evaluated to derive the recommended quality of water for various uses.

Water quality objective A numerical concentration limit or narrative statement that has been established to support and protect the designated uses of water at a specified site. It is based on scientific criteria or water quality guidelines but may be modified by other inputs such as social or political constraints.

Wastewater Water that has been adversely affected in quality by anthropogenic influence such as use in a washing, flushing, or manufacturing process.

Watertable The level of groundwater; the upper surface of the zone of saturation for underground water.

Whole effluent toxicity testing (WET testing)

See Direct toxicity assessment.

Xenobiotic A foreign chemical or material not produced in nature and not normally considered a constituent of a specified biological system. This term is usually applied to manufactured chemicals.

Zooplankton The animal portion of the plankton.

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References ANZECC/ARMCANZ (2000) Australian and New Zealand Guidelines for Fresh and Marine Water Quality.

AS/NZS 2031:2001, Selection of containers and preservation of water samples for microbiological analysis.

AS2360:1993, Measurement of fluid flow in closed conduits.

AS/NZS 2865:2001, Safe working in confined spaces.

AS/NZS 3550.7:1993, Waters. The construction and use of the Secchi disc.

AS/NZS 3778:2001, Measurement of water flow in open channels.

AS/NZS 5667.1, .4 .11:1998, Water Quality—Sampling—Guidance on the design of sampling programs, sampling techniques and the preservation and handling of samples.

AS/NZS 5667.12:1999, Water quality—Sampling—Guidance on the Sampling of Bottom Sediments.

ASTM (American Society for Testing and Materials), Annual Book of ASTM Standards. Section 11 Water and Environmental Technology, Volumes 11.01-11.02, ASTM.

AWWA (2005) American Water Works Association, Standard Methods for the Examination of Water and Wastes, 21st edition.

DEH (1998) Fish Kill Reporting and Investigation Manual, Queensland Department of Environment and Heritage, November 1998.

DNRM (2004) Queensland Harmful Algal Bloom Operational Procedures, Queensland Department of Natural Resources and Mines, February 2004.

EPA (2006) Queensland Water Quality Guidelines, The State of Queensland, Environmental Protection Agency, March 2006.

IATA Dangerous Goods Regulations, current annual edition, International Air Transport Association, Montreal and Geneva.

MDBC (1997) Murray Darling Basin Groundwater Quality Sampling Guidelines, Murray Darling Basin Commission, Technical Report No. 3, Groundwater Working Group.

NHMRC (2004) Australian Drinking Water Guidelines.

NHMRC (2006) Guidelines for Managing Risks in Recreational Water.

NRM (2001) Australia-Wide Assessment of River Health: Queensland AusRivAS Sampling and Processing Manual, Monitoring River Heath Initiative Technical Report No. 12, Commonwealth of Australia and Queensland Department of Natural Resources and Mines, Canberra and Rocklea.

QHSS (1998) Queensland Health Scientific Services, Guidelines for Nutrient Sampling of Waters, Guideline QSE ENG 001, Revision 2.

Queensland Animal Care and Protection Act 2001.

Queensland Division of Workplace Health and Safety (1991) Code of Practice on Selection, Provision and Use of Personal Protective Equipment.

Queensland Environmental Protection (Water) Policy 1997 (subordinate legislation 1997 No. 136).

Simpson, S.L., Batley, G.E., Chariton, A.A., Stauber, J.L., King, C.K., Chapman, J.C., Hyne, R.V., Gale, S.A., Roach, A.C. and Maher, W.A. (2005) Handbook for Sediment Quality Assessment (CSIRO: Bangor, NSW).

USEPA (1997 or current version) Methods for Chemical Analysis of Water and Wastes. United States Environmental Protection Agency, Publication no. EPA-600/4-79-020.

Wruck, D.J. and Ferris, J. (1997) Collection and Storage of Water Samples for Nutrient Analysis, Workshop on Sampling Nutrients in Aquatic Ecosystems: Collecting Valid and Representative Samples, University of Canberra, 21-22 April 1997.

Documents

Monitoring and Sampling Manual 2009 · Monitoring and Sampling Manual 2009 ... 3.3.2 Preservation and storage ... Appendix H7 National taxonomic codes for macroinvertebrate families