A Protocol for Responding to Adverse Fish Health Events and ...

A Protocol for Responding to Adverse Fish Health Events and Chemical 

Overspray/Drift Incidents at Sunland Fish Hatchery by Munro Mortimer 

Context 

If and when future adverse fish health events occur at the Sunland Fish hatchery, it 

may be appropriate to collect environmentally relevant materials (water, soil, 

sediment, pond vegetation, whole fish and fish tissues, surface wipes, and deployed 

droplet capture devices) for laboratory analysis. 

Capacity to respond in an effective and timely manner involves having staff trained 

in the techniques required, and having the necessary equipment on hand at an 

appropriate location/s. 

Collecting environmental samples for chemical testing 

It is essential that all samples are collected in such that: 

1. Material sent for analysis is representative of the environmental location 

sampled; 

Ensuring materials are representative usually means collecting composite 

samples. For example in sampling a pond, water samples should be taken 

from several points and mixed in equal proportions to get a composite for 

analysis. 

Unless water is well‐mixed and shallow, contaminant concentrations may 

vary with depth. This needs to be accounted for in the sampling design ‐. E.g. 

composites through depth or discrete composites at standard depths. 

The same principles apply to sampling plants, fish, soils, sediments. 

2. If comparisons are to be made between locations or against a benchmark, 

then statistical requirements should be met (e.g. multiple composite 

samples). The relevance and availability of comparable material from 

reference locations should also be considered; 

3. The interpretational requirements of the data should be built into the 

sampling design; 

For example if water is being sampled, is it a filtered sample or an unfiltered 

sample (or both) that is required? If filtration is appropriate, then field 

filtration may be necessary; 

4. Not contaminated as a result of incorrect handling or storage; 

A schedule of the appropriate containers and preservatives is attached. This 

is taken from the Qld EPA (now DERM) Water Quality Sampling Manual as 

updated for the new EPP(Water) Sampling Manual 2009. 

1

In general, amber solvent‐washed glass bottles with Teflon lid liners are 

required for organics, and acid‐washed glass or plastic (not for mercury 

analysis) bottles are required for metals and metalloids. 

Gloves should be worn and changed frequently. Ideally, samples are collected 

directly into the storage container. If this is impractical, an interim container 

may be used but care is needed to avoid cross‐contamination (e.g. clean 

between samples/sites). 

Fish tissues (or whole animals), plants/plant tissues, soils, and sediments, 

should be stored in solvent‐washed glass or foil wrapped (organics), or stored 

in acid‐washed glass or plastic for metals analysis. 

5. Loss of contaminant between sampling and analysis is minimised (covered 

under 4 above); 

Fish and fish tissues for veterinary examination 

For pathology, the procedures published in “Cultured Aquatic Animals – preparation 

for Veterinary Laboratory Examination (Second Edition)” QDPI should be followed. 

Sprayed surfaces 

Sampling of sprayed surfaces (e.g. buildings, pathways, vegetation) can be done by 

taking wipe samples. The protocol from the Qld EPA (now DERM) Water Quality 

Sampling Manual as updated for the new EPP(Water) Sampling Manual 2009 is 

attached. 

Overspray and Spray Drift sampling 

Advice on appropriate absorbent pads has been sought from UQ, Gatton. They have 

indicated willingness to provide materials and advice – a modest quote to cover cost 

of materials is expected. 

Quality Control 

Field blanks and blind replicates should be included. 

Samples should be labelled as they are collected and a written record made (what, 

where, when, by whom, etc.). Coded labels are fine (e.g. initials and site numbers 

marked on a map). 

2

Table 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, PET, polypropylene or 

similar) [‡] Except where specifically stated otherwise, put sample in container first, THEN add the preservative/s – if more than one, in the order shown thus: [A] [B]. 

[§] ‘Refrigerate’ = cool to 1‐4C; ‘Freeze’ = freeze to ‐20C. [†] Denotes recommended maximum time from collection in the field to analysis in the laboratory. 

Container Code 

Preservation Procedure 

(as above) 

Filling Method 

Rec. Max. 

Quality Characteristic 

How much: (a) OR (b) 

and/or 

Preservative 

Storage Conditions holding 

(WATER SAMPLES) V W M Sample Filtering (if any) 

to add [‡] 

(a) Mass or (b) to pH 

[§] 

period [†] 

Volume shown 

Other Instructions 

ACIDITY 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 

(A.O.X.) 

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 200 See G or 

Refrigerate Maximum 

Escherichia coli and 

note P 

24 hours 

THERMOTOLERANT 

BACTERIA 

Enterococci 

[2] 

See note [ 3 ] 

BIOCHEMICAL OXYGEN 

DEMAND (B.O.D.) 

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 None — — — 1 month — 

exclude air 

BROMIDE 100 D P — None — — Refrigerate in the dark 1 month — 

CHEMICAL OXYGEN DEMAND 100 A G or Fill container completely to Sulphuric Acid — 1‐2 Refrigerate 

7 days — 

(C.O.D.) 

P exclude air 

Freeze 

1 month 

CHLORIDE 100 D P — None — — — 1 month — 

CHLORINE, Free 100 D P — None — — Keep sample out of 5 min Determine immediately on site 

direct sunlight 

CHLORINE, Total 100 D P — None — — Keep sample out of 

direct sunlight 

5 min Determine immediately on site 

CHLOROPHYLL 

use 1 of the 2 methods 

see note [4] 

See 

note 

[4] 

1000 D G or 

P 

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 

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 — 

3

CONDUCTIVITY 


CYANIDE, Total 

see note [ 5 ] 

100 W P Fill container completely to None — — — 24 hours Determine in situ or on site if possible 

exclude air 

Refrigerate 1 month 

500 W P — Sodium Hydroxide Solution* — 12 Refrigerate in the dark 24 hours *Caution: this procedure can create 

lethal HCN gas 

DISSOLVED OXYGEN (D.O.) 300 W G — None — — — — Determine in situ or on site 

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 

HERBICIDES AND PESTICIDES: 

Carbamates 

Pyrethrins 

Synthetic pyrethroids 

Organo‐chlorine, 

Organo‐phosphorus and 

Nitrogen‐containing 

Pesticides 

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] 


see note [6] 

80 mg per L 

sample 

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 

— Refrigerate 7 days * Lid of sample container must have 


(poly‐tetrafluoroethylene) 

HYDROCARBONS, Petroleum 1000 S G Do not pre‐rinse container with Sulphuric Acid — 1‐2 Refrigerate 1 month — 

sample material. Fill bottle 

completely with sample (no 

head space) 

IODIDE 500 D P — None — — Refrigerate in the dark 1 month — 

LIGNINS AND TANNINS 250 W G — None — — Refrigerate 7 days — 

METALS – Calcium or 

Magnesium 

see note [ 7 ] re preserving; 

METALS Chromium, 

Hexavalent 

see note [ 8 ] 

250 A P Fill container completely to 

exclude air 

Nitric Acid — 1‐2 — 1 month — 

None — — — 7 days 

100 A P — None — — Refrigerate 24 hours — 

4

METALS – Mercury 250 A G For ‘Dissolved’ form, filter on 

site; see note [9] 

METALS– Potassium or Sodium 


[A] Nitric Acid — 1‐2 — 1 month If sample is wastes or contaminated 

waters, more Potassium Dichromate may 

be needed:— 

[B] Potassium Dichromate 

50 mg/mL 

Per 250 mL 

sample— 

5 mL or more 

see last 

column 

— • 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 

100 W P — None — — — 1 month — 

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] 

NUTRIENTS: AMMONIA 


NUTRIENTS: NITRATE 


NUTRIENTS: NITRITE 


250 A P For ‘Dissolved’ form, filter on 

site; see note [9] 

Nitric Acid — 1‐2 — 1 month — 

100 W P Filter on site; 

see note [9] 

None — — Refrigerate 24 hours Store in area free of contamination 

(ammonia vapour may permeate the 

Filter on site; 

see note [9] 

None — — Freeze 1 month walls of containers, even if made of high 

density polyethylene) 

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

Filter on site; 

see note [9] 

None — — Freeze 1 month 

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

after collection 

Freeze 48 hours 

5

NUTRIENTS: NITROGEN, TOTAL 

KJELDAHL (T.K.N.) or TOTAL 

(T.N.)use 1 of the 2 methods 

NUTRIENTS: PHOSPHORUS, 

Dissolved use 1 of the 2 

methods 

NUTRIENTS: PHOSPHORUS, 

Total use 1 of the 2 methods 

250 W P — None — — Refrigerate 24 hours — 

100 W P Filter on site; 

see note [9] 

Freeze 1 month 

None — — Refrigerate 24 hours — 


250 W P — None — — Refrigerate 24 hours — 

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 


(poly‐tetrafluoroethylene). Protect 

sample from light 

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

POLYCHLORINATED BIPHENYLS 

(P.C.B.) 

POLYNUCLEAR AROMATIC 

HYDROCARBONS (P.A.H.) 






1000 S G Do not pre‐rinse container with 






see note [6] 


see note [6] 

80 mg per L 

sample 

80 mg per L 

sample 

— Refrigerate in the dark 7 days * Lid of sample container must have 


(poly‐tetrafluoroethylene) 

— Refrigerate in the dark 7 days — 

SOLIDS, Dissolved 1000 D P Fill container completely to None — — Refrigerate 24 hours — 

exclude air 

SOLIDS, Suspended 1000 D P — None — — Refrigerate 24 hours — 

SULPHATE 200 W P — None — — 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 

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 

6


see note [6] 

Ammonium chloride 0.2g per 200 

mL sample 

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

16 mg per 

200 mL 

sample 

7

Table 3 ‐ Sediment 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: PTFE, polyethylene, PET, polypropylene or 

similar) [‡] Except where specifically stated otherwise, put sample in container first, THEN add the preservative/s – if more than one, in the order shown thus: [A] [B]. 

[§] ‘Refrigerate’ = cool to between 1‐4C; ‘Freeze’ = freeze to ‐20C. [†] Denotes recommended maximum time from collection in the field to analysis in the laboratory. 

Container Code 

Preservation Procedure 

Rec. Max. 

Quality Characteristic (as above) Filling Method Preservative How much: (a) OR (b) Storage Conditions holding Other Instructions 

(SEDIMENT SAMPLES) V W M to add [‡] (a) Mass or 

Volume 

HEAVY METALS 


(b) to pH 

shown 

[§] period [†] 

375 A G — None — — — 7 days — 

Ziplock Plastic Bag — None — — Freeze 1 month 

ORGANOTINS (e.g. TBT) 375 S G None — — Refrigerate 7 days — 

PESTICIDES AND HERBICIDES 


SEMI‐VOLATILE ORGANICS 

375 

375 

S 

S 

G 

G 

— 

— 

None 

None 

— 

— 

— 

— 

Refrigerate 

Refrigerate 

7 days 

7 days 

— 

— 

TOTAL ORGANIC CARBON 375 S G — None — — Refrigerate 7 days — 

(T.O.C.) 

See note [12] 

Ziplock Plastic Bag — None — — Freeze 1 month 

8

Note [1] 

Note [2]. 

Note [3]. 

Note [4]. 

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. 

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 neutralizes the chlorine, thus preventing further bactericidal effects on organisms in the 

water during transit. 

AS/NZS 2031‐2001 indicates that microbiological examination should commence within 6 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. 

Filter the sample and submit the filter paper itself (containing the particulates so removed) 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 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 (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]. 

Note [7]. 

Note [8]. 

Note [9]. 

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

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. 

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. 

Filter sample in the field, through 0.45 m poly ether sulphone filter, preferably using fully‐enclosed pressure filtering equipment, such as single‐use syringe (50 mL) with 

0.45 m 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 on the sample. 

9

Note [12]. 

If analysis is required for total organic carbon (T.O.C.) as well as for PESTICIDES AND HERBICIDES, the same sample will suffice for both analyses. 

10

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. Swipe 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 cms 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 of each 

11

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 each used swab 

sealed in a labelled sampling container appropriate for the storage of the analytes of 

interest, for example a solvent‐washed jar if pesticides are of interest. 

. 

12

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