ECASA

Book of Protocols for Field Work

Introduction

We are compiling a "toolbox" that is based on indicators of environment - aquaculture interactions (impacts) which will then be employed in predictive models. There is a need to assure the quality of field and laboratory data collected and an essential element in data quality assurance is standardization of the methods that are used to collect the data.

This compilation of protocols is a rough draft, with widely variable formats, that will serve as the basis for discussion of best methods at the January 24-25, 2006 Oban meeting. After we have selected the best protocol for measuring each variable, we will establish a uniform format and produce a "book of recommended protocols" as a deliverable of the ECASA project.

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Quality AssuranceICES Techniques in Marine Environmental Sciences, No. 32

ObjectivesThe objective of this document is to guide organizations (or individuals) towards the establishment of QA procedures, often for the first time, which will ensure that the data generated are suitable for contributing to international-level assessments of environmental quality. While some elements of any newly incorporated QA scheme must, from the outset, be considered mandatory, past experience suggests the need for a pragmatic view of how such a scheme will initially proceed. Thus, enhancements in performance may well be step-wise, in response to the adoption of new in-house working procedures, and as lessons are learned from intercomparison exercises, workshops, and other relevant activities. At this stage, a prevailing climate of encouragement will be the most helpful in facilitating such a progression.

THE QUALITY SYSTEMGeneral“Quality system” is a term used to describe measures which ensure that a laboratory fulfils the requirements for its analytical tasks on a continuing basis. A laboratory should establish andoperate a Quality System adequate for the range of activities, i.e., for the type and extent ofinvestigations, for which it has been employed. The Quality System should refer to methodology, organization and staff, equipment and quality audit (see Annex 2). The Quality System must be formalized in a Quality Manual that must be maintained and kept up-to-date. Some comments and explanations are given in this section.

Topics of Quality AssuranceIn practice, Quality Assurance applies to all aspects of analytical investigation, and includes the following principal elements:• A knowledge of the purpose of the investigation, which is essential to establish the requireddata quality.• Provision and optimization of appropriate laboratory facilities and analytical equipment.• Provision and regular updating of taxonomic keys and supporting literature for identification of biological specimens, including allowance for the possibility of the occurrence of introduced species.• Selection and training of staff for the sampling and analytical task in question.• Establishment of definitive instructions for appropriate collection, preservation, storage, and transport procedures to maintain the integrity of samples prior to analysis.• Use of suitable pre-treatment procedures prior to the analysis of samples, to prevent cross contamination and loss of the determinand in the samples.• Validation of appropriate analytical methods to ensure that measurements are of the required quality to meet the needs of the investigations.• Conduct of regular intra-laboratory checks on the accuracy of routine measurements, by the analysis of appropriate reference materials, to assess whether the analytical methods are correctly employed and remain valid. Typically, control charts are used to evaluate the findings.• Participation in inter-laboratory quality assessments (proficiency testing schemes, ring tests, training courses) to provide an independent assessment of the laboratory’s capability to produce reliable measurements.• The preparation and use of written instructions, laboratory protocols, laboratory journals, etc., so that specific analytical data can be traced to the relevant samples and vice versa.

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• The establishment of national/regional lists of all species likely to be encountered in surveys of marine communities, employing up-to-date nomenclature and recognized coding systems such as Species 2000 (see http://www.sp2000.org/), Encyclopaedia Taxonomica (see http://www.taxonomica.com/Taxonomica2/Introduction.asp) and the Integrated Taxonomic Information System (http://www.itis.usda.gov/).• The management of the information in a suitable certified database/information system.

In-house Quality ManualEvery phase of a monitoring or assessment survey, even in small laboratories, must be enforced to ensure the quality of data acquisition, collection, handling and analysis, and subsequent data management and reporting. In-house Quality Manuals must be developed in accordance withappropriate national and international standards and followed rigorously. The person responsible for authorization and compilation of the Quality Manual should be identified. A distribution list of the quality manual and identification of holders of controlled copies of the quality manual should be included. The in-house quality manual should contain, as a minimum, the following items or their equivalent:1) Scope;2) References;3) Definitions;4) Statement of quality policy;5) Organization and management;6) Quality system audit and review;7) A formal listing of the staff involved in the monitoring, analytical, and technical work as well as quality control management with respective training, professional qualification, and responsibilities within the laboratory;8) Standard Operating Procedures (SOPs) (see Section 2.3.1, below);9) Certificates and reports;10) Sub-contracting of calibration or testing;11) Outside support services and supplies;12) Handling of complaints;13) Contingency planning for the eventuality of problems arising (see also Section 4.1, below).

Standard Operating Procedures (SOPs)An SOP may be defined as “a documented procedure which describes how to perform tests or activities normally not specified in detail in study plans or test guidelines” (Good Laboratory Practice, 1997). The italicized text helps to clarify some confusion that exists with regard to the role of an SOP. For example, in cases where international guidelines for a sampling or analytical procedure are written in sufficient detail, then these will perform the same function. However, guideline documents frequently cover large sea areas and a variety of habitats and cannot be expected to provide sufficient detail for the requirements of all local surveys. In these circumstances, an SOP bridges the gap between the activity of an individual laboratory and the wider need for harmonization of methodology. For example, a laboratory SOP might include a description of sample-processing equipment peculiar to that laboratory (though compatible with the performance needs of external guidelines), and perhaps its local source of manufacture. A well-written SOP will help inexperienced members of staff in a laboratory to quickly develop expertise in a sampling or analytical area which is consistent with past practice at that laboratory, while being compatible with established approaches elsewhere. For those seeking laboratory accreditation, the production of SOPs will be essential as part of a

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wider QA package but, even for those who are not, they provide an important means to foster good practiceinternally. However, SOPs are clearly not, in themselves, guarantors of data quality. SOPs should describe all steps performed in biological measurement. They should be established to cover the following areas of activity:• Station selection and location, navigational accuracy;• Handling, maintenance, and calibration of field and laboratory equipment;• Handling and use of chemicals (i.e., fixatives, preservatives, reagents) used in marine environmental surveys;• Collection of biological material;• Storage of biological material including labelling, and checking of preservation status;• Distribution of biological material to external contractors/taxonomic specialists;• Analytical methods for biological material;• Identification of biological material including taxonomic expertise of the personnel;• Recording of biological and environmental data and subsequent data management;• Analysis of biological and environmental data;• QA of report writing and documentation including signed protocols in all steps of analysis. SOPs should contain a description of operational procedures. An outline structure for an SOP (modified from ISO/IEC, 1999) is as follows:• scope of procedure used;• description of the study target;• variable to be determined;• equipment necessary, reference material (e.g., voucher specimens) and taxonomic literature used;• specification of working conditions required for effective sampling;• description of procedure/method with respect to the following aspects:i) sampling and sample treatment, labelling, handling, transport and storage of samples, preparation for laboratory analysis,ii) instrument control and calibration,iii) recording of data,iv) safety aspects;• criteria to adopt or reject results/measurements;• data to be recorded and methods for their analysis;• assessment of uncertainty of measurements.In considering “best practice”, it is recommended that SOPs should:• be structured logically by heading and sub-heading to cover the full sequence of activities in field sampling and laboratory analysis;• carry an issue number, date, and name(s) of the individual(s) responsible for its drafting and updating. This anticipates a likely requirement for changes to SOPs in response to new equipment, guidelines, and so on;• document in-house AQC procedures;• account for the specific practices of the individual laboratory. At the same time, SOPs must of course reflect agreed guidelines applicable at national or international level, for example, relating to nomenclature and coding systems employed in documenting the outcome of the analysis of field-collected specimens;• contain a full listing of taxonomic keys used for laboratory identification, and other useful reference works relating to procedures;• be filed as paper copies in an accessible place, as well as being available on a computer network;• be freely available to all interested parties (especially funding agencies);

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• contain explicit instructions for the tracking of samples from the point of collection to the point of archiving of analysed material. SOPs may usefully contain:• diagrams depicting gear, especially where local modifications to equipment are made;• a summary flow-chart as an accompaniment to a lengthy SOP, as an aide memoire for field and laboratory bench operators;• details of local suppliers, manufacturers, etc., where relevant.SOPs should not:• contain vague generalizations;• contain excessive detail: a sensible balance needs to be achieved which takes into account the basic level of training and common sense that a new operator will possess;• cover too many activities: for example, it is logical to have separate SOPs for field and laboratory procedures. Different types of field activity such as intertidal core sampling and ship-board sampling are also sensibly treated separately.ConclusionThe preparation of SOPs to cover field and laboratory analytical activities is one of the most important practical steps that a laboratory/institute can take in seeking to improve the quality and consistency of its scientific output and is, therefore, to be strongly recommended. This having been done, inter-laboratory comparisons of SOPs may then provide a useful tool in identifying any remaining inconsistencies, and hence in promoting harmonization of methodology at a national and international level. Such periodic comparisons of SOPs are also to be strongly recommended (see, for example, Cooper and Rees, 2002).

Organization, Management, and StaffOrganizationThe Quality System should provide general information on the identity and legal status of the laboratory and should include a statement of the technical role of the laboratory (e.g., employedin marine environmental monitoring). The information must include general lines of responsibility within the laboratory (including the relationship between management, technical operations, quality control and support services, and any parent or sister organizations). In the case of smaller units, the organizational tasks must be allotted to fewer personnel or even one individual.ManagementClear job descriptions, qualifications, training, and experience are necessary for all persons concerned with QA and QC. Job descriptions should include a brief summary of function, the pathways of reporting key tasks that the jobholder performs in the laboratory, and limits of authority and responsibility.StaffMinimum levels of qualification and experience necessary for the engagement of staff and their assignment to respective duties must be defined. Members of staff authorized to use particularitems of equipment should be identified and the institution should ensure that all staff receive training adequate to the competent performance of the relevant methods and operation of equipment. A record should be maintained which provides evidence that individual members of staff have been adequately trained and that their competence to carry out specific methods, identifications or techniques has been assessed. Managers should be aware that a change of experienced and well-trained staff might jeopardize continuity in the production of data of consistent quality. In the case of small units employing few staff or even single individuals responsible for the generation of data, a scheme for the certification of individual expertise

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(e.g., in aspects of species identification) may be a valid alternative to formal accreditation involving a hierarchy of quality managers, which may not be practicable.

EquipmentAs part of its quality system, a laboratory is required to operate a programme for the necessary maintenance and calibration of equipment used in the field and in the laboratory to ensure against bias of results. General service equipment should be maintained by appropriate cleaning and operational checks, where necessary. Calibrations will be necessary where the equipment can significantly affect the analytical result. Performance checks and service should be carried out at specific intervals on microscopes, balances, and other instruments. The frequency of such performance checks will be determined by experience and based on the need, type, and previous performance of the equipment.

DocumentationAll biological data produced by a laboratory should be completely documented (“meta-information”) and should be traceable back to its origin. The necessary documentation should contain a description of sampling equipment and procedures, reference to SOPs for the sampling, sample handling and analytical procedures involved, and the names of persons responsible for Quality Control. In general, one signed protocol should accompany a sample through all steps of processing.

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Selection of sampling stationsMERAMED project field handbook Once the hydrographical features have been determined for the study site (see below), sampling stations may be determined on the basis of current direction and intensity. At some sites, there may be seasonal differences in dominant current direction and velocity and these should be taken into consideration when planning where to sample.

Using the information from current meters or profilers, a transect will be established along the prevailing direction of the water currents. Samples will be taken at a station directly below the edge of the cages (station "0") and at 25 and 50m from the edge of the cages along the transect line. A control station will be chosen 1 km upstream from the cages at similar depth and sediment type. In the event that a 1 km reference station is not feasible (this happens in some cases), a different station will be used. To establish station "0" one either stands on walkway and deploys sampling gear from there or from a boat tied up next to the cages. Other stations are distance from the cage edge (0m station). A station below the center of the farm (cages) is not sampled, because even if divers are available as this station it becomes unusable in future surveys if divers are not available.

Note: In the eastern Mediterranean, stations were established 0, 5, 10, 25 and 50m downstream from the edge of the cages. This may not apply to salmon or other forms of aquaculture and/or non-Mediterranean environments. Partners may use additional stations depending on the "dispersiveness" of their environment. For depositional environments, partners should add 5 and 10 m stations, for deep dispersive environments partners should add 75 m or AZE edge stations (see below).

Current regulatory trends regarding station selection in Scotland are:- transects perpendicular to main axis of flow not relevant- a farmer gets an Allowable Zone of Effect, within which the main considerations are

the sediment health at cage edge and the extent of the boundary of the AZE and the conditions at the boundary; conditions in the transitional zone of perpendicular to the main current is supplementary (unnecessary information)

- 0, 25 and 50 m in the direction of the residual current (downstream), with 25 and 50 m upstream (i.e. 5 stations)

- sample at cage edge and 10 m either side of the boundary of the AZE. The AZE boundary is determined from information available such as the modelled situation for the maximum biomass, previous surveys (i.e. 3 stations). Identification/checking of the AZE boundary to some extent may also be undertaken on site with visual assessments or quick measures with sulphide/Redox probes

- dispersive sites may have a station at a distance greater than 50 m; this may be the case for deep sites where the footprint of initial deposition covers wide are, but near-bed current is low and so resuspension is minimal

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HydrographyMERAMED project field handbook Hydrography is one of the key features used in site selection: a) flux of good quality water via the finfish or shellfish gear (water quality, animal welfare); b) spatial dispersal of effluents, which may spread load over sufficiently large area to reduce benthic impact on seafloor (environmental health), c) supply of planktonic food to shellfish (enhanced growth). Hydrographic characterization needs to be done well before the intensive WP5 site study as it will serve as the basis for pre-selecting sampling stations (see below).

The most common types of instruments used for hydrographic measurements around fish farms are rotary, electromagnetic and acoustic doppler current profilers (ADCP). The first two instrument types measure current speed and direction at a discrete depth, the third type profiles the water column with some limitations at surface and bed.

A sampling interval of 10 or 20 minutes is recommended for long-term deployments (i.e. up to 1 month). Short-term deployments (i.e. of a few days) should use a 5-minute sampling interval. Where acoustic profilers are being used, care should be taken to ensure that the standard deviation of the measurements is sufficiently small. Total battery life and memory capacity should be taken into account when setting up the instrument.

Select a mooring type depending on local site conditions (i.e. level of exposure of site, local fishing activities), intended mooring location, instrument type and length of deployment. The mooring should be located outside the shadowing effect of the cages. U-shaped mooring arrangement is recommended for long-term deployments as it allows three methods for recovery in the event of the surface buoys being lost. Where a surface marker is lost, recovery can take place from the opposite end. Where both surface markers are lost, recovery can take place by snagging the groundline with a grapnel.

A conductivity, temperature and depth (CTD) profile is useful at the deployment and recovery stages of the hydrographic instruments. This gives information on the depth of any water column features in relation to instrument depth. The depth and length of deployment of each instrument depends on the objectives of the current meter study. It also relates to any modelling intended for the survey site, in which case the modelling guidelines can be consulted.

The following recommendations are made with respect to current meter deployment depth:*if three instruments are available – deploy surface, mid-water and near-bed *if one instrument is available – deploy mid-water or below the depth of the cage bottom*if two instruments are available, then consideration should be given to the level of resuspension expected at the site before deciding on instrument deployment depths. Near-bed current speed exceeding 10 cm s-1 causes resupension of fish farm material and so near-bed measurements are crucial at sites where these conditions exist. Therefore, deploy surface and near-bed if greater than 5 % of near-bed current speeds are expected to exceed 10 cm s-1, otherwise deploy surface and mid–water.If profiler is available – deploy either ‘looking down’ or ‘looking up’ but consideration must be given to the effect of surrounding fish farm equipment and the limitations of current measurement at the surface and near-bed

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Actual deployment depths will depend on the total depth of the water column, but measurement of surface current and current below the bottom of the net pen is desirable. In the case where a counter current to the surface flows exists at depth, this should be measured if possible.

A deployment of 1 month is recommended for studies aiming at measuring the general hydrodynamic features of a site. The effect of season must also be taken into account where possible, as seasonal wind patterns are likely to change hydrographic patterns. Measurement of current in the same season as any intensive benthic sampling is ideal, but this may not be practical. Need to select one sampling station which should avoid shadowing effect of cages but should be sufficiently close to cages to sample general flow conditions in the area of expected deposition.

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A. Water Column Variables

Secchi Disk Transparency Protocol Water clarity is a measurement of how far down light penetrates into the water column.This indicates the amount of suspended particles such as plankton and/or silt in thewater.Need: Secchi disk (weighted) with rope marked to 5cm resolution . ClothespinsIn the Field

1. Record the cloud cover (see Cloud Cover Field Guide below). 2. Stand so that the sun is to your back (to avoid glare) and so it will be easy to see the

Secchi disk where the measurement will be made. 3. If you cannot reach the water surface, establish a reference height. This can be a railing,

a person’s hip, or the edge of a dock. All measurements should be taken from this point. 4. Remove sunglasses. Lower the disk slowly into the water until it just disappears. 5. Mark the rope with a clothespin at the water surface or at the reference height. 6. Raise the disk until it reappears and mark the rope with a clothespin at the water surface

or at your reference height. 7. The Secchi depth is the average of these two depths. You may wish to repeat this test yourself or have a second viewer replicate the measurement.

8. Record depth of Secchi disk disappearance and reappearance, time of day, tide,current condition (strong, mild, or none), wind/wave condition (ripples, small waves, whitecaps), viewer’s initials and any other significant notes.

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Chlorophyll a - extractive methodPlymouth Marine Laboratory Sampling1. Filter e.g. 1000 ml water sample through 47mm (or 25mm) GF/F filter at relatively low vacuum pressure (<250 mm Hg). Smaller volumes may be adequate, e.g. 10 to 100ml depending upon predicted chlorophyll concentration.2. Fold filter in half (sample side inwards) place in lidded petri-dish and freeze until required.

CHLOROPHYLL EXTRACTIONRequired: spatula, flat forceps, 10ml pipette and measuring cylinder, 90% acetone, homogeniser and stand, homogeniser tube, centrifuge tubes and lids, acetone wash bottle and discard pot. 1. For natural seawater and more delicate species (e.g Phaeocystis; Isochrysis ) : place filter in bottom of centrifuge tube, add 10 ml 90% acetone then refridgerate (4°C) for between 4 and 24 hours 1. Gently invert tube several times before decanting off into fluorometer tube or spectrophotometer cuvette (see further on for fluorometer or spectrophotometer protocols).2. For more robust species: place filter at the bottom of homogeniser, tube then add 2 - 4 ml taken from a measured volume of 10 ml of 90% Acetone. Homogenise at low speed for approximately 20 seconds.a) Pour into centrifuge tube, rinse homogeniser tube with remaining 6-8 ml 90% acetone then add to that already collected in centrifuge tube, scrape out any remaining bits with spatula and add these to centrifuge tube also. Cap centrifuge tube and store in refrigerator for between 4 and 24 hours. Clean off homogeniser head and tube between samples.b) Remove samples from fridge and centrifuge for 5 mins at 3,000 to 4,000 rpm (if centrifuge has cooling facility then set this at 5 - 10 °C).

Throughout storage and analysis of samples, exposure to light (especially strong sunlight) should be avoided or at least minimised.

FLUOROMETER DETERMINATION (using Turner Designs fluorometer)2

Require: acetone wash bottle (90% acetone), pasteur pipette, 1N HCL, discard pot, tissues,

1. Switch on fluorometer (approx. 30 mins prior to use) and ensure it is in MANUAL mode2. Insert a blank (i.e 90% acetone) should read zero.3. Insert sample and establish required range i.e x 1 or x 100 (using black knobs at side of machine) depending on concentration of chlorophyll in sample4. Put range onto AUTO (it will then automatically click up through the ranges, x30, x10 etc, and select the most suitable one. Read off the scale value (as 0 to 100 for both x1 and x100 ranges) against the given FA value on the calibration table provided (this should be stuck on top of the Turner in the basement). You only need the FA valuee.g for a range x1 x30 the FA value = 5.9 (µg litre)e.g for a range x100 x30 the FA value = 0.06 (µg litre)

1 Note that time filters left in fridge MUST be standardised; assumption is that up to 95% of the chlorophyll is released in the first 4 hours, but the remainder is released very slowly, expect c. 100% recovery after 24 hours. If experiment involves making comparisons between robust/non-robust species then ALL filters must be homogenised.2 See Strickland J.D.Hand Parsons T.R. A Practical Handbook of Seawater Analysis. Fisheries Research Board of Canada (Ottawa 1968) Bulletin 167.

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5. Add 100 µl (2 drops from pasteur pipette) of 1N HCL and read off. Should the auto range differ from that selected prior to the addition of HCL go into MANUAL mode and select the original range. Chlorophyll.a (µg litre) = (FA (RB - RA)*Ve (ml))/Vf (ml)

FA = calibration factor of fluorometer (this was calculated as µg litre)RB = fluorescence reading before addition of 1N HCLRA = fluorescence reading after addition 1N HCLVe = Vol.of acetone extract (ml)Vf = Vol. of sample filtered (ml)

Calculation Example (Fluormetric)a sample of 200 ml was filtered and 10 ml 90% acetone was used for extraction, the scale was read off on x 1 x 30 (hence FA=5.9); RB=49 ; RA=31.75 ; hence chl.a = 5.08875 µg litre

SPECTROPHOTOMETER DETERMINATIONRequire : 2 matched cuvettes (4 cm path length), glass rod for stirring HCL, acetone wash bottle, pasteur pipette, discard pot, tissues, 1N HCL.

1. Prepare as for fluorometry.2. Switch on spectrophotometer 30 mins prior to use. If not using sipper - set switch inside to normal not remote.3. Set wavelengths to 750 and 663 and bandwidth to 1 nm. Ensure digital display knob is set on abs.4. on programme controller panel:a) turn button onb) wavelength mode to SELECTc) programme mode to SINGLEd) turn wavelength control knob to 750 nm, then move white line of lander 1 switch around to the front until red light is triggered one) turn wavelength control knob to 663 nm, then move white line of lander 4 to front until red light comes on (nb landers 2 and 3 should be set on c. 100 nm if not required, using same process as abovef) then turn wavelength mode to FIXED.5. SETTING UP CELLS:a) Fill both cuvettes with 90% acetone and place one in farthest cell holder (= reference cell), and the other in front cell. Read off to ensure that they are both paired (should read approx. 0.0) if not then swap cuvettes around, should then be ok.b) remove cuvette from front cell, fill with sample, switch to 750 nm, then zero, press blue manual override button and go down to 663 nm and read off abs. Return to 750 nm (via red manual override button) add 2 drops 1N HCL (pasteur pipette) and leave c.30 secs. Rezero if nec. , go down to 663 nm and read off abs.c) rinse cuvette with 90% acetone between samples.

Chlorophyll a (µg litre) =( 26.7((E663 - E750) - (E663a - E750)) x vol.extract (ml)) / (vol.filtered (litres) x path length (cm) )

E663 = abs. prior to HCLE663a = abs. after addition of HCL

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Quality AssuranceSampling QA• Keep the samples cool and in the dark.• For chlorophyll a it is recommended that the sample is filtered immediately after samplingor, at least, as soon as possible thereafter; avoid deposition of cells. If storage isunavoidable, the filters should be deep frozen (< -20 oC).Spectrophotometric or fluorometric chlorophyll a analysis QA • The analysis should follow ISO 10260; departure from this has to be documented, andevidence of comparability of the data provided.• The samples/filters and the chlorophyll a extracts should be handled in subdued light.• Avoid evaporation of the extraction solvent during extraction and measurement procedures.• The measurements should be done immediately after clearing the extracts; the preference isfor equipment for measuring the whole spectrum (800–350 nm) for easier checking ofshifting of the chlorophyll peaks.• Validate the spectrophotometer and the fluorometer at least once a year, or when changesof the equipment are required.• Calibrate the equipment with a certified reference material, if possible; use control charts.

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NUTRIENTS (N, P) in seawaterICES Techniques in Marine Environmental Sciences, No. 35 64

The commonly designated nutrients are inorganic nitrogen compounds (NO3 -, NO2 - NH4+),phosphate (PO4 3-) and silicate (SiO4 3-). Total phosphorus (Ptot) and total nitrogen (Ntot) are also included because of their importance in relation to ecosystem analysis and budgets. Nutrients in sea water are considered trace determinands and their analysis is liable to various sources of contamination. Sea water for nutrient analysis is usually collected from research vessels or ships of opportunity (e.g., ferry boats, fishing boats, coast guard or navy vessels). The reference method for measuring nutrients in the Baltic Sea (including storage and pre-treatment) is Grasshoff (1976) “Methods of Seawater Analysis”.

SAMPLE HANDLINGSpecial attention must be paid to possible nutrient sample contamination generated by the ship. Wastewater discharged from wash basins, showers, and toilets contains significant amounts of phosphorus and nitrogen compounds and, therefore, can contaminate the surface waters to be sampled. For this reason, the water sampler must be deployed far from wastewater outlets, even if no sewage is discharged at the time of sampling. Although most modern ships are equipped with special sewage tanks, they are often emptied at sea owing to a lack of appropriate reception facilities in ports. In addition, there are potential problems with kitchen garbage. Mixing by the ship’s propeller can disturb the natural distribution of the determinands in the surface layer, particularly as regards oxygen. These problems, including the exact location of the ship, should be considered along with the natural variability. Phosphorus and nitrogen compounds are secreted from human skin. However, touching of the sampler and the sample bottles by hands does not cause problems unless the sample comes into contact with the outer surface of the sampler or sample bottle. This is something that should never happen since the outer surfaces cannot be kept free of contamination on-board a ship. In view of the potential for contamination, the analyst should preferably supervise the collection of samples. The attaching of bottles to a hydrowire or the preparation of a rosette and the subsequent removal and transport of samples to the ship’s laboratory should be done by trained personnel.The written instructions for the collection of samples should include the precautions to be taken when a sub-sample is transferred to the storage container. The instructions must include the details of the essential record of the sample: station location, station code, depth of sampling, date, time, etc., and the identity of the person responsible for sampling.

STORAGE OF SAMPLESThe stability of nutrients in seawater samples depends strongly on the season and the location from which the samples were taken. Nutrients in seawater samples are generally unstable. Grasshoff (1976) recommends that ammonia and nitrite are measured no later than one hour after sampling. Samples for nitrate, phosphate, and silicate should preferably be analysed within six hours after sampling, and no later than ten hours. If for practical reasons samples cannot be analysed within these time limits, the corresponding data should be flagged if stored in databases, unless the storage method has been validated. Samples should be stored protected from light and refrigerated. Plastic bottles must be used if silicate is measured. New sample bottles sometimes adsorb nutrients onto their walls. The newbottles, if necessary, should be cleaned with phosphate-free detergent, rinsed generously with distilled/deionized water, and left filled with sea water containing nutrients for a few days. Then checks for adsorption of nutrients onto the walls or losses due to transformation to another chemical form should be carried out. Sample bottles should always be rinsed with the

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seawater sample from the sampler before they are filled. As regards ammonia determination, glassware for ammonia should always be cleaned with dilute hydrochloric acid. If samples cannot be analysed within the above-mentioned time limits, the following methods of storage can be recommended.Silicate - 0–4° C protected from light. Do not freeze (polymerization may occur).Nitrite - Freezing or 0–4° C protected from light. Do not acidify (rapid decomposition). Ammonia - No known preservation methods are applicable.Nitrate - Freezing.Total nitrogen - Freezing or 0–4° C protected from light. Do not acidify (enhanced risk of contamination).Phosphate - Freezing or acidification.Total phosphorus - Freezing or acidification with sulphuric acid, store at 0–4° C, protected from light.

The addition of mercury or chloroform is an alternative preservation method for all nutrients except ammonia. However, these chemicals can affect the reaction kinetics, especially with automated methods, and this effect should be evaluated by the laboratory. The same chemical preservation of calibrants and quality controls can compensate for this effect. The use of mercury should be minimized and optimum disposal procedures should be ensured. These preservation methods are all second choice to immediate analysis. They should, as mentioned, be validated by each laboratory, taking into account the concentration levels, storage time and environment, differences in sample matrices, and the analytical method of the laboratory. Since no preservation method for nutrients can, at present, be recommended for general use, each laboratory must validate its storage methods for each nutrient before they are used routinely.

SAMPLE PRE-TREATMENTSea water contains microorganisms and other suspended matter of different composition. In some cases, these particles bias the measurement of the determinand in the soluble phase. The suspended matter can be removed either by filtration or centrifugation. Unnecessary manipulation of the sample should be avoided, but in particle-rich waters (e.g., coastal waters, during plankton blooms), filtration or centrifugation may become necessary. It is important that the procedure used for filtration/centrifugation has been validated.For removing algae from the water sample, a GF/C filter is adequate. For work in open oceans with low concentrations of suspended matter, GF/F filters are considered suitable for suspended matter separation from open sea water. Filtration in closed systems with a neutral gas is recommended. Centrifugation is especially advisable for samples destined for ammonia determination. If a sample containing particles is not filtered, the turbidity causes light scattering which can bias a colorimetric measurement. In this case, a turbidity blank should be carried out by measuring light absorption of the sample before adding the colour-forming reagents.

APPROPRIATE CHEMICAL ANALYTICAL METHODSThe choice of an analytical method should be based on the following criteria:• the method should measure the desired constituent, i.e., be adequately specific, with accuracy sufficient to meet the data needs in the presence of interferences normally encountered in natural samples;• the method should be sufficiently simple and rapid to permit routine use for the examinationof large numbers of samples.

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The reference methods used for manual nutrient measurements are described by Grasshoff (1976). Any changes to the reference methodology should be validated before use for routine work. Apart from manual methods, various automated methods are in use, including different types of continuous flow analysis (CFA, steady-state mode, and peak mode) or flow injection analysis (FIA or Reverse Flow Injection). The analyst has to be aware of the effects of the different analytical conditions in automated analysis which might affect accuracy.

CALIBRATION AND THE BLANKStock standard solutions should be prepared separately for each determinand using analytical grade reagents that can be pre-treated to a precise stochiometric composition, e.g., by drying excess moisture. Reagents containing crystal water should be dried at a sufficiently low temperature in order not to remove the crystal water (the drying temperature is compound dependent). Stock standard solutions containing more than 1 mM are stable for long periods (up to one year refrigerated), but working calibration solutions must be prepared daily and used within hours of preparation.Blank sea water may be prepared from a bulk sample of offshore surface sea water collected in summer, when the nutrients are at low or below-detection concentrations (Kirkwood, 1994). Blank sea water and reagents totally devoid of nutrients are, however, difficult to achieve, especially regarding the content of ammonia. Optimum handling precautions should be taken to minimize the content of nutrients to below approximately 10% of the measuring range. The concentrations of nutrients in the blank and reagents can be assessed by the standard addition method.For ammonia analysis, the salinity of the samples affects the reaction kinetics, mainly due to the buffer effect of marine water which results in a sub-optimum end pH. This effect can give biased results, especially with kinetically dependent automated methods. In the Baltic Sea, the salinity ranges from approximately 0 to 30, and therefore the size of this bias will be variable. This kinetic effect should be checked by standard addition, or by checking the pH of the reagent-sample mixture, which should be in the range between 10.5 and 11. Whenever compensation for this bias is deemed necessary, one of the following methods is suggested:a) If all samples have the same salinity, calibrate using the addition of calibrants to one of the samples. In some situations, low-nutrient sea water can be prepared by aging and filtering natural sea water (as mentioned above).b) Empirical correction in accordance with the measured sample salinity or pH value. For all photometric nutrient measurements, differences in light refraction, caused by differences in the salt concentration, can give rise to shifts in blank/baseline values, especially in light-measuring cells with round windows. This can be compensated by using blanks and calibrants of the same salt concentration as the samples.Particles can give rise to light-scattering effects that result in interferences in all photometricnutrient analyses. This bias can be avoided by measuring the sample before addition of thecolour reagent, or by filtration or centrifugation where this does not cause contamination.

REFERENCESGrasshoff, K. 1976. Methods of seawater analysis. Verlag Chemie, Weinheim, New York. Kirkwood, D. 1994. Nutrients: Practical notes on their determination in seawater. In ICES/HELCOM Workshop on Quality Assurance of Chemical Analytical Procedures for the Baltic Monitoring Programme. Ed. by G. Topping and U. Harms. Baltic Sea Environment Proceedings No. 58: 23–47.

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Suspended Particulate Matter (SPM = total particulate matter, TPM), Particulate Inorganic Matter (PIM), Particulate Organic Matter (POM) Plymouth Marine LaboratoryRequired - pre-washed, ashed and weighed GF/F 47mm filters prepared as below, stored in clean petri slides, clean forceps, 100% acetone in wash bottle.

Filter-preparation:a. To remove fine loose particles of filter, separate and soak in distilled water for > 1h; agitate and rinse 3-4 times in distilled water.b. Partially dry each filter on suction head to remove excess water (this prevents sticking to foil in the next step).c. Place filters individually into foil envelope/fan and oven dry overnight.d. Carefully number each filter on the exposed margin (soft lead pencil or pre-tested pen) and lay out (slightly overlapping) on foil tray, fit a lid and ash in muffle furnace at 450°C for >4h.e. Cool in dessicator; all handling of filters, from this point on, using clean (acetone) forceps only to avoid contamination.f. Remove individually and weigh to 5 places, standardising the time it takes to weigh (filters increase in weight as they take up atmopheric moisture), and store place in numbered petri-slides.

Processing of filters for measures of POM and TPM is as follows:a. Filter through required volume of homogenised material.b. Rinse with 0.5M Ammonium formate (31.5 g/l) to remove salt; 10 ml twice through filtered sample and also with water bottle around margin having removed filter head (all with pump running), and return to petrislide.c. Oven dry (60°C for 2 days, 40°C for 1 week).d. Weigh (from dessicator to 5 places, as above, preferably with the same balance) for total particulates (TPM).e. Ash at 450°C in muffle furnace for > 4hf. Weigh (from dessicator to 5 places, as above, preferably with the same balance) for for inorganic particulates (PIM).g. Do all of the above using at least 10 blank filters (prepared and processed as above, but without sample) for each experimental day (changes in weight before and after experimentation are used to correct for changes in balance calibration and/or filter water content).

Absolute care in the preparation and processing of these filters as described is essential, for small errors in weight at these stages will significantly bias ratios and other results calculated later. Many experiments have been ruined by lack of attention to the above details!

3. CHN - 25mm filters prepared as below, stored in acid-washed petrislides.Necessary to remove all organics and keep free from contamination using gloves and covers etc.

Filter-preparation is as follows:a. Ash at 450°C for >4h, laid out in foil tray with foil cover.b. Cool in dessicator.c. Store in box until required.

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d. When using: filter material through, then wave filter in HCl vapour for 15 secs to destroy inorganic material (do not rinse filter), then place in numbered acid-washed petri-slide and oven dry at < 40 oC.

e. Upon removal from oven, seal petri-slide with cover.

1 SAMPLE PROCESSING

The amount of sample (volume of water) required will depend on water quality. Measures of Chl a are very sensitive, so that one only needs enough sample on the filter such that one can see a change in colour. Measures of CHN require about 4 times more sample volume on filters of the same size.

However, the measure of POM and TPM requires as much sample as is reasonable on the filter, and certainly more than 2 mg TPM. Especially when filtering natural seawater, this means filtering until the filter is almost blocked. This may only require 500 mls in times of algal bloom or resuspension, but when seston levels are low, we have filtered up to 3 litres per sample (UK waters).

All filtering must be quantitative (i.e. we need to know the initial total volume of each sample, and the separate volumes of that sample filtered for separate determinations of Chl a, CHN and POM/TPM; thus allowing us to calculate the total of each within the sample as a whole). Use the data sheets supplied to help ensure this. Examples under conditions of average food availability are as follows:

1.1 Inflow seston ( = outflow from control chamber without animal, which represents food availability)

1 litre bottle (shake well)

for TPM, PIM, POM for CHN for Chl a 300ml on 47mm GF/F 50ml on 25mm GF/F 10ml on 25mm GF/F (rinse with NH4 formate) (acid fumes) (no rinse)2 duplicate filters 2 duplicate filters 2 duplicate filtersTherefore, for each sample, require: 2 47mm GFFs (washed, ashed, weighed)

2 25mm GFFs (ashed only)2 25mm GFFs (no pretreatment)

Faeces ( = true or pseudo, analysed to calculate rejection and absorption efficiencies)

Each sample may be adjusted to a standard volume (say 22 ml) using filtered seawater. From this:

1 animal: 22ml (mix well)

for TPM, PIM, POM for CHN for Chl a 15ml on 47mm GFF 2ml on 25mm GFF 0.5ml on 25mm GFF (rinse with NH4 formate) (acid fumes) (no rinse)1 filter only 1 filter only 1 filter only

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Therefore, for each sample of either true or pseudofaeces, we require:1x 47mm GFFs (washed, ashed, weighed)1x 25mm GFFs (ashed only)1x 25mm GFFs (no pretreatment)

After filtering, place all CHN and POM/TPM filters in the low temperature (< 40 oC) oven to dry (more than 40 oC will result in loss of lipids). Chl a samples must not be dried; instead, fold in half with material on inner side, and stored in freezer.

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PARTICULATE ORGANIC CARBON IN SEA WATERICES Techniques in Marine Environmental Sciences, No. 35 147

INTRODUCTIONParticulate matterThe particle size of the organically bound carbon of particles (POC) generally ranges between 0.45 µm and 300 µm. This includes both living organisms, such as phytoplankton, yeasts, bacteria, and microzooplankton, and detrital particles and aggregates. The production and decomposition of biogenic particles as well as their fractional removal to the deep sea control the distribution of most trace elements in the oceans. Microbial decomposition, desorption, and dissolution of suspended or sinking marine particles can release elements associated with labile (e.g., organic) fractions back to the sea water. On the other hand, particles can scavenge trace elements from the dissolved phase and thereby transport them to sediments. Analysis of the composition and distribution of the particulate fractions in the oceans is therefore required to understand the behaviour and geochemical cycling of, e.g., trace elements.

Dissolved MatterAmong the different carbon reservoirs, dissolved organic matter (DOM) has the greatest mass, representing about 1000 × 1015 g of carbon, and not least because of its importance for the global climate is there a need to obtain accurate and comparable data on dissolved organic carbon (DOC) concentrations. Methods for the determination of DOC developed at a rather slow pace due to difficulties related to the composition of sea water. While DOC concentrations are around 1 mg dm–3, sea water usually contains more than 35 g dm–3 of salts and more than 25 mg dm–3 of inorganic carbon as CO2, HCO3 -, and CO32-.

SAMPLE HANDLINGThe sample should be handled and transferred between containers as little as possible to avoidcontamination during the steps between sampling and analysis (see Grasshoff et al., 1999 and ISO, 1999). It is important to obtain a representative sample, which under certain circumstances, e.g., during heavy algal blooms, can be achieved by shaking the water sampler immediately before taking the sub-sample. The homogeneity of the sample may be verified, for example, by separately analysing sub-samples from the upper and lower layers of the bottle. For POC determinations, suspended particles are collected on filters. Since organic carbon is to be measured, filters must be made of inorganic material, e.g., glass fibre or metal foil (precombusted for 4 hours at 450 °C). Whatman GF/F glass fibre filters are recommended. The determination of DOC implies that the samples are filtered. The limit between dissolved and particulate organic carbon is determined by the filter porosity (generally 0.45 µm). If the water samples are not filtered, the organic carbon content analysed would represent TOC, i.e., the sum of organically bound carbon present in water, bonded to dissolved or suspended matter.

STORAGE OF SAMPLESFilters containing particulate matter collected for POC analysis should be dried under vacuum for at least one day and stored dry in a desiccator with silica gel or, alternatively, temporarily stored in a freezer and later dried in a drying oven at 60 °C for 30 min.

APPROPRIATE CHEMICAL ANALYTICAL METHODSFor POC analysis, a variety of similar instruments currently appear on the market. In particular, Carlo Erba and Hewlett-Packard CHN analysers have frequently been used. The

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main components of the analysers are basically the same, with an autosampler, a combustion column reactor, a reduction column, a gas chromatographic separation system, the detector unit, and an output device for the analytical results. Helium is used as the carrier gas. In the combustion reactor, oxygen gas and other oxidizing and catalysing reagents support the completeness of high-temperature combustion of organic carbon and nitrogen compounds to carbon dioxide, elemental nitrogen, and nitrogen oxides. Elemental copper in the reduction column reduces nitrogen oxides to N2 and binds excess oxygen. Water and the combustion products CO2 and N2 are separated by gas chromatography, and N2 and CO2 are detected and quantified by thermal conductivity detectors (TCD).

CALIBRATION AND THE BLANKThe analysis of POC is most often carried out together with the analysis of PON (particulate organic nitrogen). For POC and PON determinations, the instrument is calibrated with high purity acetanilide (analytical grade reagent). Acetanilide is used because its elemental composition matches the elemental composition of particulate material obtained from sea water, i.e., C:N = 8. At least ten filters should be analysed to determine the procedural blanks and the standard deviations from the mean values. These filters are treated in the same way as the sample filters, but the same water which is used for rinsing the sample filters (filtered sea water or artificial sea water) is filtered through the blank filters.

INTERNAL QUALITY ASSURANCE AND CONTROLThe internal quality control should be carried out to check the operational performance of the system, by regularly analysing control samples and duplicate samples. If acetanilide is used as a control sample for POC and PON, it should be from another batch and preferably bought from another company than the calibration standard. The control samples should be analysed with each series of samples and duplicate samples should be analysed regularly. These results should be plotted on control charts in order to verify the accuracy of the results, and estimate the measurement uncertainty.

REFERENCESGrasshoff, K., Kremling, K., and Ehrhardt, M. (eds.) 1999. Methods of seawater analysis. VCH, Weinheim, New York.ISO. 1999. Water quality – Guidelines for the determination of total organic carbon (TOC) and dissolved organic carbon (DOC). ISO 8245. International Organization for Standardization, Geneva.

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BIOASSAY STUDIES WITH PHYTOPLANKTONUNIVERSITY OF CRETE

The following methodology is a modification of the protocol described by Dalsgaard (2005)

1. BackgroundRelease of nutrients from fish farms is traditionally monitored by analyzing dissolved nutrients in the waters around the fish cages. Two major drawbacks appear with this approach: the release of nutrients varies diurnally suggesting a sampling around the clock for documenting this release, which would lead to a large number of samples and a high cost for monitoring; the nutrients lost from fish cages are diluted in large volumes of water rendering the documenting of their small increase in concentration difficult by standard analytical techniques. The use of bioassays can overcome the above mentioned problems as it integrates the effects of the aquaculture over time and responds to all bio available nutrients, organic or inorganic. The approach of bioassay is simply to expose phytoplankton or macroalgae to the waters next to the aquaculture facility for a period of 3-6 days and measure the growth of these primary producers as a function of distance from the facility and thus describe the horizontal extent of the effects of nutrient release.

2. Equipment and materialsSurface water Collected from the control station of the site25 m mesh sieve/plankton netSpectra/por 1 dialysis membrane Consisting of regenerated cellulose with a

molecular weight cut off of 6-8 kilo Dalton. The flat width of the membrane is 10 cm

Plastic coated metal wireNylon mesh bagsMetal rod/plateRopeWeights OptionalBuoysAnchors

2. Bioassay setup1. Cut the dialysis membrane into pieces of 30 cm length2. Soak the pieces of the membrane in distilled water until they become soft (1-2h)3. Close one end of them with a plastic coated metal wire4. Filter surface water from the control station of the site through a 25 m mesh

sieve to remove larger grazers5. Dispense the filtered water into the dialysis bags (ca 600 ml/dialysis bag)6. Close the dialysis bags with plastic coated metal wire7. Five replicate bags will be needed for each station (ca 6.4 cm diameter, ca 20 cm

long)8. Place each bag in a nylon mesh bag for hanging and protecting the dialysis bags9. Using a rope and a metal plate hold together 5 replicate bags (Fig. 1)

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buoy

mooring weight

dialysis bagsweight (optional)

metal plate

10. A buoy on top and an anchor at the bottom will be needed for holding each bioassay setup ca 1.5 m below the surface at each site (Fig. 1)

11. At each station one bioassay setup should be incubated for 5 days.

Figure 1. Bioassay setup. Designed by the IMBC team participating in MedVeg project (based on Dalsgaard 2005).

3. Analysis 1. Biomass of phytoplankton is measured as chlorophyll-a concentrations according to

standard protocols used for water column measurements2. Before the analysis the volume of each dialysis bag must be recorded

4. Literature Dalsgaard T (2005) Bioassays with macroalgae and phytoplankton: tools for monitoring

nutrient release from fish farms. In: Effects of nutrient release from Mediterranean fish farms on benthic vegetation in coastal ecosystems Final report, p 22-31

Mura MP, Agusti S (1996) Growth rates of diatoms from coastal Antarctic waters estimated by in situ dialysis incubation. Marine Ecology Progress Series 144: 237-245

Mura MP, Agusti S, delGiorgio PA, Gasol JM, Vaque D, Duarte CM (1996) Loss-controlled phytoplankton production in nutrient-poor littoral waters of the NW Mediterranean: In situ experimental evidence. Marine Ecology Progress Series 130: 213-219

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Sediment geochemistry and benthosbased on: www.ifm.uni-kiel.de/fb/fb3/ex/sbb/heye/bequalm/framesets/Bequalm.htm & Field operations manual for marine water-column, benthic, and trawl monitoring in Southern California (www.sccwrp.org/tools/methods.htm#4-1)

Sediment sampling

Sampling Field Log should include:– person responsible for sampling;– project or contract identification code;– sketch of sampling station transect in relation to cage groups– cage group numbering system (so that biomass for cages can be matched to husbandry

sheets)– geographical co-ordinates for each sampling station (for each replicate sample in case

of boat drift during sampling), including time of day, whether the ship was anchored or not, weather conditions during sampling;

– water depth (m) at each sampling station;– sampling programme for each sampling station (number of samples, sampling of

background parameters etc.);– sampling device used for each station and each replicate sample– date sampling station;– other comments (such as rejected samples, delays and any problems experienced and

the causes);– quality of sample i.e. disturbed/not disturbed;– List of items to be reported can be found at - http://www.ices.dk/env/repforlindex.htm.

Benthic sampling using (e.g.) a Van Veen grabA 0.1 m2 modified Van Veen grab may be used to collect sediment samples for physical, chemical, and infaunal analysis (Figure 2) (Stubbs et al. 1987). The grab may be galvanized, stainless steel, or Teflon-coated. All surfaces of the grab must be clean and free of rust. Either single or tandem Van Veen grabs are acceptable. Grabs will be used if:* a large (surface area and volume) sample is required, e.g. for macrofauna* it is not possible to sample with cores by divers or using corer* however - grabs are not suitable for surface layer sampling (e.g. for meiofauna or some other analyses) because they disturb and "blow off" the surface layer

Grab sampling proceduresPrior to deployment, the grab is cocked with the safety key in place. The grab is then hoisted over the side, the safety key is removed, and the grab is lowered at 2 m/sec until it is 5 m above the bottom. From this point, it is lowered at 1 m/sec to minimize the effects of bow wave disturbance. After bottom contact has been made (indicated by slack in the lowering wire), the tension on the wire is slowly increased, causing the lever arms to close the grab. Once the grab is back on board, the top doors are opened for inspection.

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Criteria for acceptable grab samplesUpon retrieval of the grab the acceptability of the sample must be determined. Acceptability is based upon two characteristics of the sample: sample condition and depth of penetration. Sample condition is judged using criteria for surface disturbance, leakage, canting, and washing (Figure 3). Acceptable sample condition is characterized by an even surface, with minimal surface disturbance, and little or no leakage of the overlying water. Heavily canted samples are unacceptable. Samples with a large amount of "humping" along the midline of the grab indicating washing of the sample during retrieval are also unacceptable. While some humping will be evident in samples from firm bottoms where penetration has been poor, this is due to the closing action of the grab and is not evidence of unacceptable washing.

Figure 2. Modified Van Veen grab sampler recommended for marine recieiving-water monitoring programs in Southern California: a) cocked position; b) tripped position (modified from Stubbs et al. 1987)

Figure 3. Examples of acceptable and unacceptable grab sample condition

If the sample condition is acceptable, the overlying water is drained off and the depth of penetration determined. The overlying water in grabs intended for infaunal samples may be drained but all drained water must be captured for screening with the sediments (see Sample Processing below). Extra caution should be taken to drain the overlying water from the grabs for chemistry and toxicity samples. It is recommended that siphoning or

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decanting be employed for these grabs to avoid disturbance and loss of the surface sediments. It is important to get the best sample possible. For infaunal samples, sediment penetration depth must be at least 5 cm; however, penetration depths of 7-10 cm should be obtainable in silt (fine sand to clay). The depth of penetration is determined by insertion of a plastic ruler vertically along the grab midline and measurement of the depth of sediment to the nearest 0.5 cm.

Sediment DescriptionThe field description of sediments is required following measurement of penetration depth. The sediment description should encompass the following:

at minimum the sediment should be characterized as being shell hash, gravel, sand, or mud (silt and/or clay), but if possible, the following should be done:

surface colour and colour change with depth as a possible indicator of redox state; smell: sulfide (the odor of H2S or rotten eggs), oily (the odor of petroleum tar), or

humic (a musty, organic odor). Typically, sediments will have no particular odor. General sediment colors (e.g., black, green, brown, red, yellow) description of sediment types, including important notes, e.g., the occurrence of

concretions, loose algae, etc. When describing the sediment, the recommendations presented on the ICES web page should be followed. The use of stainless steel buckets or box corers is advocated in cases where the sediments are to be sub-sampled for trace metal and organic contaminants determinations. It is recommended that measurements of redox potential and shear strength be made in samples collected by a box corer rather than a grab because the latter has a great chance of distorting the sample.Precise position fixing during sampling is essential. The position and the depth should be controlled and documented by track plotting during station work.

Sediments may also be sampled using cores.a) Diver-sampled cores.Requirements:i) sediment depth must not exceed 30mii) divers need to be experienced at both taking sediment cores correctly & efficiently and they must have prior experience with the local sediments (removing cores from sandy sediments is not the same as removing cores from clayey sediments).iii) suitable cores must be made (I would prefer that all partners use same size (diameter) cores to minimize artifacts due to different wall effects on the contained sediments, especially since we'll have no control over how long a time will pass between sediment sampling and sample processing; similar material - acrylic, plexiglass, PVC, perspex - plastic and transparent) and stoppers obtainediv) carrying baskets are essential to allow divers to sample and swim with cores held upright in vertical position throughout

b) Gravity cores.these will be used, if available, instead of diver-sampled cores, if:

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* seafloor is too deep for divers* use of divers is too complicated or expensive* there's a need for core as opposed to grab samples

requirements:* gravity corer (e.g. Craib-corer )* boat with winch

Sediment geochemistry

Sediments should initially be analyzed for: a) simple measure of grain size distribution (φ-scale: silt/clay fraction < 63 µm, 125 µm, 250 µm, 500 µm, 1000 µm, 2000 µm); b) median grain size for the upper 5 cm.Additional variables of interest, for sediment indicators, as established by the sediment geochemistry sub-group are: Sediment carbon quality (ind# 1), Sulphide/oxygen with probes/total sulphur (ind# 3), Redox (ind# 11), Total N (ind# 12), Total P (ind# 13), Total organic carbon (ind# 14, 15)

Practically-speaking, need to decide how vertical profiles will be measured? Wildish (2004) proposes using cores with holes predrilled in a spiral pattern to facilitate porewater removal by syringe. Holes are covered by tape to enable sampling. In this approach care needs to be exercised not to draw down porewater from other depths (determined by the volume being removed). Another option is to measure vertical profiles using needle-type electrodes, which entails mounting electrodes on a stand and introducing probes with micromanipulator into the sediment? At what resolution should such profiles be made? For redox and hydrogen sulfide, special care is needed to avoid oxygenating the cores on one hand and letting the closed volume of water/sediment respire the contained oxygen to a state of hypoxia/anoxia (artifacts related to the time the sample is left waiting for analysis after removal from the seafloor)

Replicates: in general, need 3 replicates from each station and depth for total N, total P, organic C; LOI.

Sediment redox potential, Eh (mV)

1. Background The oxidation-reduction (redox) conditions in the surficial sediment depend on the degree of organic enrichment and can be assessed by measuring the vertical redox potential profile in the top 15 cm (expressed in mV) (Zobell, 1946). The redox state of sediment is the result of a combined effect of biological and chemical processes of reversibile and/or irreversibile nature and, therefore, difficult to define. It has been pointed out that the concentrations of the components actually reacting reversibly are, in most cases, too small to get reliable results from redox measurements. Thus what really is measured is a mixed potential which is not useful for chemical equilibrium calculations (Bågander, 1978). However, Eh profiles still provide useful indications, since the decrease in Eh with

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the depth is related to the decrease in the dissolved oxygen concentration in the pore water. Negative Eh values are therefore associated with anoxic conditions, in which the degradation of the organic matter is carried out by anaerobic bacteria. In marine sediment, these bacteria use mainly sulphate as electron acceptor which is reduced to hydrogen sulphide. (Porrello et al., 2005; Chamberlain, 2002; Aleffi et al., submitted; Danovaro et al., 2004).

2. Equipment and materials

Box corer (15x17x26,5) cm Surface sediment 0-5 cm (5 cm surface layer, collected from the sediment sample)Falcon test-tube Broad knifeMeter (to measure sediment height)Portable pH/Ion meter (specifications described below)Redox electrode with Platinum ring indicator (specifications described below)

2. Performing a pH measurement and Analysis

Electrode calibration: The ZoBell’s is used as reference in the checking routine. This solution (0.003 M potassium ferricyanide, 0.003 M potassium ferrocyanide, and 0.1 M potassium chloride) has an Eh value of +430 mV at 25°C.

Measurement:The measurement is performed in situ, by means of a portable pH/ion meter with a combination electrode. The electrode is placed in the sample and an automatic or manual endpointing (which determines the end of each single reading) is selected for the measurement. Between each measurement the electrode is rinsed with MilliQ water and wiped with a soft paper tissue.

3a. pH/ion meter specification model: portable SevenGo SG8- pH-meter/ion measurement technique specification:

3b. Electrode specification

pH mV Measurement range from -2 to 19.999 from -1999.9 to+1999.9 Resolution 0.001 0.1 Accuracy ±0.002 ± 0.1

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A combination electrode for measuring the oxidation/reduction power of a solution. Consists of a platinum ring indicator electrode and a silver/silver chloride reference electrode. The measuring signal is produced on the surface of the precious metal by an exchange of electrons with the oxidation-reduction system of the measured medium.

Specifications Temperature range °C 0 ... 80 Metal Platinum Shaft material Glass Type of frit Ceramic Reference system ARGENTHAL, Ag+ trap Reference electrolyte 3mol/l KCI Shaft length 120 mm Shaft diameter 12 mm

4. LiteratureAleffi, I.F., Bettoso, N., Solis-Weiss, V., Tamberlich, F., Predonzani, S., Fonda-Umani, S., submitted to ICES – Journal of Marine Science. Effects of suspended mussel culture on the macrozoobenthos in the Gulf of Trieste (Northern Adriatic Sea, Italy).Chamberlain, J., 2002. Modelling the environmental Impacts of Suspended Mussel (Mytilus edulis L.) Farming. Ph-D Thesis, Napier Univeristy, Edimburgh.Danovaro, R., Gambi, C., Luna, G.M., Mirto, S., 2004. Sustainable impact of mussel farming in the Adriatic Sea (Mediterranean Sea): evidence from biochemical, microbial and meiofaunal indicators. Marine Pollution Bulletin, 49: 325-333.Lars Erik Bågander, 1978. An evaluation of the use of redox measurements for characterizing recent sediments. Estuarine and coastal Marine Science, 6: 127-134.Porrello, S., Tomassetti, P., Manzueto, L., Finoia, M.G., Persia, E., Mercatali, I., Stipa, P., in press. The influence of marine cages on the sediment chemistry in the Western Mediterranean Sea. Aquac.Zobell, C. E., 1946. Studies on redox potential of marine sediments. Bulletin of the American Association of Petroleum Geologists 30, 477-511.

Organic carbon determination in surficial (1 cm) sediment

1. Background

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1.2 Carbon in marine sediment occurs as organic carbon intimately linked to the metabolic processes of plants and animals and as carbon contained within biogenetic and abiogenetic carbonate minerals. Successful determination of organic carbon relies upon the separation of organic from inorganic carbon. Several methods of analyses have been employed for the separation of organic from inorganic forms of carbon (Verardo et al.,1990). Although the determination of total carbon (TC) content in marine sediments is straightforward by a carbon-hydrogen-nitrogen (CHN) elemental analyzer, that of organic carbon (OC) presents difficulties. These because organic material is a complex mixture, and some of these components may be lost from the sediment at the temperatures both above and below those for the loss of other materials, such as structural water and carbonates. The determination of OC is usually a lengthy procedure, involving weighing of the sediment sample, careful pretreatment of the sample by a suitable acid to remove IC, drying of the sediment, weighing of sediment and analysis of the sediment by CHN analyzer (Tung & Tanner, 2003).

2. Equipment and materialsBox corer (15x17x26,5) cm (to sample surface sediments)Falcon test-tube Broad knifeMeter (to measure sediments height)Centrifuge (to extract interstitial water from the sediments)Vacuum freeze dryer (to lyophilize the sediments)Mortar (to omogenize sediments)Sieve with mesh of 200 µmMicro-analytical balance Portable refrigerator (to store sediments during sampling)

2. Determination of organic carbon in sediments

Sampling: Collect the sediment sample, by means a Box-corer device; Store in a Falcon test-tube the upper 1 cm layer of the sediment sample;

Sample processing: Sediment sample is placed into a portable refrigerator and then transported to the

laboratory; Samples are centrifuged to remove interstitial water; Samples are stored at –20°C until analysis; A freeze-dryer system is used to dry the wet sediment (the process can take up to a

few days); Samples are sieved at 200 µm; 6-10 mg of sediment are taken for the analysis (weighted in a soft Silver container); Treat each sample by adding 20 µl of HCl 1N. If effervescence continues, this step of

the acidification procedure must be repeated; Wait for 10 minutes;

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Exsiccate samples in oven, drying for 10 minutes at 60°C; Addition of 20 µl of HCl 1:1. If effervescence continues, this step of the

acidification procedure must be repeated; Wait for 10 minutes; Exsiccate samples in oven, drying for 10 minutes at 60°C; Addition of 20 µl of HCl 25%. If effervescence continues, this step of the

acidification procedure must be repeated; Wait for some minutes; Exsiccate samples in oven, drying for one night at 60°C;

Analysis:OC concentration is determined using a carbon-hydrogen-nitrogen (CHN) Fisons Mod. EA1108 Elemental. Reference material BCSS (NRC, Canada) is adopted to assess the accuracy of the analytical data. 4. LiteratureDavid J. Verardo, 1990. Determination of organic carbon and nitrogen in marine sediments using the Carlo Erba NA-1500 Analyzer. Deep-Sea Research 37 (1): 157-165.Tung J. W. T. and Tanner, P. A., 2003. Instrumental determination of organic carbon in marine sediments, Marine Chemistry, 80 (2-3): 161-170.

Total nitrogen determination in surficial (1 cm) sediment

1. Background The production of biodeposits (faeces and pseudofaeces) due to mussel cultivation, can cause an increase in the TN concentrations in the sediment underneath mussel lines (Chamberlain, 2002; Aleffi et al., in press). In a recent EI study (Christensen et al., 2003), an higher C/N ratio in surficial sediment underneath the lines with respect to a reference site was observed.

2. Equipment and materialsBox corer (15x17x26,5) cm (to sample surface sediments)Falcon test-tube Broad knifeMeter (to measure sediments height)Centrifuge (to extract interstitial water from the sediments)Vacuum freeze dryer (to lyophilize the sediments)Mortar (to homogenize sediments)Sieve with mesh of 200 µmMicro-analytical balance Portable refrigerator (to store sediments during sampling)

3. Determination of total nitrogen in sedimentsSampling: Collect the sediment sample by means of the Box-corer device;

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Store in a Falcon test-tube the upper 1 cm layer of the sediment sample;

Sample processing: Sediment sample is placed into a portable refrigerator and then transported to the

laboratory; Samples are centrifuged to remove interstitial water; Samples are stored at –20°C until analysis; A freeze-dryer system is used to dry the wet sediment (the process can take up to a

few days); Samples are sieved at 200 µm; 6-10 mg of sediment are taken for the analysis (weighted in a soft Silver container);Analysis:OC concentration is determined using a carbon-hydrogen-nitrogen (CHN) Fisons Mod. EA1108 Elemental. Reference material BCSS (NRC, Canada) is adopted to assess the accuracy of the analytical data.

4. Literature Aleffi, I.F., Bettoso, N., Solis-Weiss, V., Tamberlich, F., Predonzani, S., Fonda-Umani, S., submitted to ICES – Journal of Marine Science. Effects of suspended mussel culture on the macrozoobenthos in the Gulf of Trieste (Northern Adriatic Sea, Italy).Chamberlain, J., 2002. Modelling the environmental Impacts of Suspended Mussel (Mytilus edulis L.) Farming. Ph-D Thesis, Napier Univeristy, Edimburgh.Christensen, P.B., Glud, R.N., Dalsgaard, T., Gillespie, P., 2003. Impacts of longline mussel farming on oxygen and nitrogen dynamics and biological communities of coastal sediments. Aquac. 218: 567-588.Ryba S. A., 2002. Effects of sample preparation on the measurement of organic carbon, hydrogen, nitrogen, sulfur, and oxygen concentrations in marine sediments. Chemosphere, 48: 139-147.

Total phosphorous determination in surficial (1 cm) sediment

1. Background The production of biodeposits (faeces and pseudofaeces) due to mussel cultivation, can cause an enrichement of organic matter in the sediment. If we also consider mussel and epibiota drop-off during harvesting, much higher rates would be expected and longline mussel production may thereby dramatically alter the benthic environment. However, relatively little is known of the effects of longline mussel farming on benthic ecology, microbial mineralization and nutrient dynamics (Christensen et al., 2003). In the aim of a better understanding of nutrient dynamics, Total Phosphorous concentration, a parameter complementary to the reactive phosphorus, can be reasonably included in a set of benthic geochemical indicators. The method here applied for the extraction of total phosphorous is the one proposed from Aspila (1976).

2. Equipment and materials Box corer (15x17x26,5) cm (to sample surface sediments)

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Falcon test-tube Broad knifeMeter (to measure sediments height)Centrifuge (to extract interstitial water from the sediments)Vacuum freeze dryer (to lyophilize the sediments)Mortar (to omogenize sediments)Sieve with mesh of 200 µmMicro-analytical balance Portable refrigerator (to store sediments during sampling)

2. Determination of total phosphorous in sedimentsSampling: Collect the sediment sample by means of the Box-corer device; Store in a Falcon test-tube the upper 1 cm layer of the sediment sample;

Sample processing: Sediment sample is placed into a portable refrigerator and then transported to the

laboratory; Samples are centrifuged to remove interstitial water; Samples are stored at –20°C until analysis; A freeze-dryer system is used to dry the wet sediment (the process can take up to a

few days); Samples are sieved at 200 µm; About 100 mg of sediment are taken for the analysis (weighted using a weighing

boat); Total phosphorus is extracted from sediments with 1 N HCl after ignition at hight

temperature (550°C for a 4 hours time) ; The dried sediment is transferred in the test-tube (the weighing boat is rinsed with 10

mL HCl 1N); Test-tube are sonificated for 15 min. and then stirred for 16 hours at ambient

temperature; Test-tubes are entrifuged; The surnatant is removed and put in a 100 mL volumetric flask; The sediment is washed with 10 mL HCl 1 N; Test-tube are sonificated for 15 min; Test-tube are stirred for 10 min; Centrifuge the test-tubes; The surnatant is removed and put in the 100 mL volumetric flask; The washing procedure is repeated again, with 10 mL HCl 1 N; The volumetric flask, containing the surnatant, is filled with HCl 1N;

Analysis:TP concentration is determined using an inductively coupled plasma atomic emission spectrometer (Spectro Modula) at 177.500 nm wavelength

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3. Literature Aspila, K.I., Agemian, H., Chau, A.S.Y., 1976. A semiautomatedmethod for the determination of inorganic, organic and total phosphate in sediments. Analyst, 101, 187– 197.Christensen, P.B., Glud, R.N., Dalsgaard, T., Gillespie, P., 2003. Impacts of longline mussel farming on oxygen and nitrogen dynamics and biological communities of coastal sediments. Aquac. 218: 567-588.

Determination of total sulphur in sediments ICRAM

For total sulfur, we collect the surface layer sediment (1.5 cm) samples using plastic corer tubes (50 ml capacity) directly by Van Veen grab equipped with screen doors. In the laboratory we homogenize by grinding all the sample collected and after, for the analysis, we use a subsample of about 10 mg of sediment - see belowThe CHNS-O EA 1110 ThermoElectron is an instrument dedicated to the simultaneous determination of amount (%) of carbon, hydrogen, nitrogen and sulphur present in a wide range of organic and inorganic substances in liquid, solid or gaseous samples. The process is known as Dinamic Flash combustion. The principle of operation is founded on three sequential steps: the sample, kept in a lightweigth tin capsule, is energetically oxidized yielding a gas mixture which is swept into a chromatographic column from which any eluted pure combustion gas passes through a thermoconductivity detector which generates an electrical output signal proportional to the amount of eluted gas. The resulting four components are detected in sequence N2, CO2, H2O, SO2.For the determination of total sulphur on marine sediment, we use a specific configuration of the instrument; in fact is need to increase the instrumental sensibility for the low sulphur concentration in marine sediment.The reaction tube is composed by 5 cm of quartz wool, 14 cm of electrolitic copper (90 gr), 1 cm of quartz wool and 5 cm of copper (II) oxide wires. The reaction tube temperature is 900°C.The gas chromatographic column, specific for the total sulphur determination, is made in Poropack PQS and the length is 0.8 mt. The oven temperature is 65°C.In order to amplify the signal is necessary set the gain at x10.In a lightweight tin capsule, about 15 mg of sediment (dried and homogenized) are weighted and added with about 10 mg of catalyst Vanadium pentoxide (V2O5), in order to ensure complete conversion of inorganic sulphur in SO2. The time for the analysis is 280 seconds.For the calibration curve we use Marine Sediment Reference Material (MESS-2), the value of sulphur is certified (0.18±0.04) by the National Research Council of Canada.

Determination of hydrogen sulphideICES Techniques in Marine Environmental Sciences, No. 35 62

Introduction

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Hydrogen sulphide is a poisonous gas that readily dissolves in water. Hydrogen sulphide is formed in stagnant waters, where the oxygen has been consumed by bacteria oxidizing organic matter to carbon dioxide, water, and inorganic ions. Sulphate-reducing bacteria then use the oxygen bound in sulphate ions as an electron acceptor while reducing the sulphate ions to sulphide. No higher life forms can exist in water containing hydrogen sulphide, and these areas are thus turned into oceanic deserts. Hydrogen sulphide in a water sample is easily detected by its characteristic smell, even at concentrations lower than those measurable with the method below.

MethodsThe reference method for sampling and the determination of hydrogen sulphide in the Baltic area is the spectrophotometric method described in Fonselius et al. (1999). This book should be consulted for exact reagent compositions and procedures. For concentrations up to approximately 250 µM, the method by Fonselius et al. (1999) is recommended. Samples with higher concentrations can be treated in two different ways. Samples containing higher concentrations may be diluted after precipitation with a zinc acetate solution containing 2 g l-1 of gelatin (Grasshoff and Chan, 1971). This solution can be homogenized and diluted. However, higher levels of sulphide are better quantified using the method by Cline (1969).

SamplingSamples are taken from ordinary hydrocast bottles immediately after the oxygen samples have been taken, using the same sampling technique (cf. Technical Note on the Determination of Dissolved Oxygen, Attachment 2, above). If no oxygen is present, the sulphide samples should be taken first. Sulphide reacts with many metals, and the samplers should thus preferably be all plastic. 50–100 ml oxygen bottles are recommended. The two reagents are added simultaneously using piston pipettes or dispensers. The tips of the pipetting devices should be close to the bottom of the bottle. No air bubbles should be trapped in the bottle. Note that the amounts of reagents added have to be adjusted according to the size of the bottles used. As concentrations rather than amounts are measured, no exact knowledge of the bottle volume is required. Samples that cannot be analysed within 48 hours may be preserved with zinc acetate, which precipitates the sulphide as zinc sulphide. The preserved samples can be stored for a few months, if light and temperature changes are avoided. Prior to analysis, the reagents are added in the same way as for unpreserved samples. When the bottle is turned, the precipitate dissolveseasily, and the colour develops normally.

4 Analytical proceduresAbsorbances are measured in a spectrophotometer or a filter photometer at 670 nm.Measurements should be performed no sooner than 1 hour and no later than 48 hours after the reagent addition.

5 Quality assuranceThe following QA elements must be satisfied:

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1) The performance of the photometer with regard to absorbance and wavelength correctness must be checked and documented using a certified set of filters, or by an equivalent method.2) The reagents must be calibrated using the procedure described in Fonselius et al. (1999). For measuring volumes in this procedure, only calibrated or class A glassware should be used. It is essential that the working solutions are freshly prepared, and that the sulphide content of the stock solution is measured, not calculated from the weighing of Na2S (as Na2S of sufficient purity is not available).3) New reagents should be prepared at one-year intervals. The old reagents always must be checked against the newly prepared reagents in order to prove their stability.4) No stable solutions are available for control charts. The difference between double samples in a control chart with zero as the reference line provides information on both precision and the validity of the sub-sampling. Ideally, the result (Sample 1 – Sample 2) should be evenly distributed around zero. Any deviations from this suggest sub-sampling problems.

6 Reporting resultsThe concentration of hydrogen sulphide is usually expressed as µmol l-1 (µM), or in some casesas ml l-1 H2S or as negative oxygen.X µmol l-1 S2- = X × 22.41 × 10-3 ml l-1 H2SY ml l-1 H2S = Y × 103 / 22.41 µmol l-1 S2-Z µmol l-1 S2- = -0.044 × Z negative oxygen units (ml l-1)

7 PrecisionUsing the method recommended in Fonselius et al. (1999), the analytical precision will be approximately ±1 µmol l-1.

8 ReferencesCline, J.D. 1969. Spectrophotometric determination of hydrogen sulfide in natural waters.Limnology and Oceanography, 14: 454–458.Fonselius, S., Dyrssen, D., and Yhlen, B. 1999. Determination of hydrogen sulphide. InMethods of seawater analysis, 3rd edition. Ed. by K. Grasshoff et al. Wiley-VCH,Germany.Grasshoff, K., and Chan, K.M. 1971. An automatic method for the determination of hydrogen sulphide in natural waters. Analytica Chimica Acta, 53: 442–445.

Protocols for Macrobenthos

Separation of Fauna from the SedimentThe transfer of the sample to the sieve, the sieving procedure, and the transfer of the animals to the fixation jar are the steps during sample treatment most likely to introduce sources of error. To reduce the magnitude of these errors, the number of steps in the sampling and sieving procedures should be kept as small as possible and attention should be paid to the following procedures.

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Sieving can be conducted either aboard the survey vessel as samples are collected or onshore after a sampling excursion has been completed. In the first case, sieving usually precedes fixation and is conducted primarily on live organisms. In the second case, sieving generally occurs after fixation and is therefore conducted on dead organisms. Comparability between the results of these two techniques may be influenced by at least two factors. First, because fixation may cause some taxa to distort their shape or autotomize (i.e., cast off body parts), the sieving characteristics of those taxa may change following fixation. Second, sieving characteristics of live organisms may differ from those of dead individuals. This bias occurs primarily for soft-bodied organisms (e.g., polychaetes) that can crawl through mesh openings or entangle themselves on the screen when they are sieved live.A major problem that may be encountered when organisms are fixed in sediment before being sieved is that the fixative either will not reach all buried organisms or will not reach them in time or in sufficient concentration to prevent some deterioration. Because deteriorated individuals may decompose completely or fragment upon sieving, their sieving characteristics can be modified substantially by inadequate fixation. Therefore, if samples are fixed in sediment, extra care should be taken to ensure that organisms are fixed adequately. For example, the sample container can be rotated gently immediately after fixation and again after 12-24 h to ensure adequate fixative penetration.From a logistical standpoint, sieving of samples in the field is generally preferred for surveys in which a large number of samples are collected during each cruise. Field sieving results in a considerable reduction in the volume of material that must be stored on the vessel (i.e., where space is often limiting) and later transported to the laboratory.

Use of RelaxantsRelaxants are often used when processing benthic macroinvertebrate samples for at least two major reasons. First, relaxants facilitate taxonomic identifications (and morphometric measurements) by reducing the tendency of organisms to distort then shape or autotomize when exposed to a fixative (Gosner 1971). Complete organisms having a natural appearance are easier to identify correctly than are fragmented and/or distorted specimens. For some taxonomic groups (e.g., Maldanidae), complete organisms are required for species-level identification.A second reason for using a relaxant is to ensure that animals are sieved whole, if sieving follows fixation. The tendency for some taxa (especially polychaetes) to autotomize if not relaxed can influence sieving by reducing the size of individuals.Because relaxation can influence taxonomic identification and sieving, data comparability between studies that use a relaxant and those that do not use one may be affected. The magnitude of these effects is unknown, but probably is greatest for soft-bodied taxa that are difficult to identify (e.g., some polychaetes) and smallest for taxa encased in a hard enclosure such as a calcareous shell (e.g., most molluscs) or an exoskeleton (e.g., crustaceans), particularly if the hard parts are the primary taxonomic characters used for identification.

Sieving optionsThere are new designs of sieving tables with hand-controlled water sprinklers, which help to reduce the physical stress on the people involved while at the same time retaining the

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quality of the sampled specimens (Figure 3). Also, tilting devices for the full sample container, providing the option to fix the container at a certain angle over the sieve, are of use to reduce spilling and to avoid destructive tools. One example of a smaller sieve holder is shown in Figure 4. With this stand, the sieve residue can be transferred to the sample container with only the help of a sprinkler bottle, thereby avoiding the need for spoons or other scraping tools.For descriptive surveys, sieves used for extraction of the macrofauna from sediments should have a mesh size of 1.0 mm. The use of an finer sieve of mesh size 0.5 mm, or even finer, is recommended for special purposes. The sieve mesh should be checked from time to time for damage and wear. If a finer sieve is also used, the sieve fractions should be treated separately, and the results should be given for the single and the summed fractions. If re-sieving of samples is carried out, a mesh size finer than that of the initial sieve should always be used. Small sieves may be cleaned with an ultrasonic bath. The use of brushes should be avoided to prevent possible alterations of the mesh size. Distortion of woven mesh sieves occurs with increasing frequency of use. This can introduce considerable errors in the collection of small organisms. Moreover, the use of a square mesh introduces additional inaccuracies in collecting organisms in the size range of approximately the mesh size since the mesh diagonal width is greater than the nominal mesh width. The use of larger sieves is encouraged because the risk of clogging is reduced, for example, sandy samples may rapidly fill or even overfill smaller sieves. Larger sieves also reduce the risk of spilling when transferring samples from containers/buckets to the sieve. This risk can also be kept low by using integrated sieve tables, as shown in Figure 3.

Figure 3. Cross-section of an integrated sieving table where the sample is first emptied onto a coarse sieve (~5 mm) from where it is washed with a hand sprinkler douche onto the final 1 mm (0.5 mm) sieve (Design provided by G. Fallesen, Aarhus, Denmark). A growing number of institutes are changing to round mesh sieves, owing partly to a perceived improvement in the condition of the animals retained and partly to the theoretical improvement in mesh selectivity. Further work is required to establish a basis for using either type of sieve. Errors associated with the use of different sieves are like to

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be small in relation to other sources of sampling error.

Figure 4. Sieve holder to provide a careful transfer of the sieve residue to the sample vessel (no tools needed-only a funnel and a wash bottle)(Design provided by G. Fallesen, Aarhus, Denmark).

Sieving ProcedureSieving should be conducted according to the following procedure:Each grab and box core sample should be sieved, stored, and documented separately. The grab or box core should be emptied into a container or washing table, and then the sample should be transferred portion by portion onto the sieves, as a sediment-water suspension. The use of sprinklers or hand-operated douches to suspend the sample is recommended. Very stiff clay can be gently fragmented by hand in the water of the container. The sieve must be cleaned after each portion has been sieved to avoid clogging and to ensure an equal mesh size throughout the entire sieving procedure. In order to avoid damaging fragile animals, the most gentle way to sieve a sample is to gently agitate the sieve surface under the water surface of a water-filled container until all sediment that can pass the sieve is washed through. On no account should water jets (i.e., deck hose) be used against the sieve surface. Fragile animals, such as some polychaetes, should be picked out by hand during the sieving, to minimize damage. Also, stones and large shells should be picked out, to avoid a grinding effect on the organisms and the sieve. All material retained on the sieve should be carefully flushed off the sieve, with water from below, into an appropriate recipient and fixed. The use of spoons or other scraping tools should be avoided. When the 0.5 mm sieve is used, the 0.5 mm and the 1 mm fractions must be kept separate throughout all further processing.

Fixation

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Fixation and conservation (preservation) are two different steps in the treatment of a sample. The former procedure is employed to coagulate and harden the tissue of the organisms, while the latter prevents them from rotting and decaying. Improperly fixed specimens may create problems during further treatment, i.e., through fragmentation of specimens or loss of appendages. Some zoological museums will only accept properly (formalin-) fixed specimens for further analysis and curation.All the material retained on the sieves should be fixed in a buffered 4 % formaldehyde solution (1 part 40 % formaldehyde solution and 9 parts filtered sea water). For buffering, 100 g of hexamethylene tetramine (= Hexamine, = Urotropine) can be used per 1 litre of concentrated formaldehyde (36-40 %). Sodium tetraborate (= Borax) in excess may also be used. Sponges are best preserved by putting them directly into absolute ethyl alcohol so as to prevent fragmentation.Formaldehyde is regarded as a toxic compound, and probably also carcinogenic, and should, therefore, be handled with great care. Appropriate means of laboratory air suction or ventilation should be provided for all procedures. For animal sorting, the samples should first be thoroughly washed with tap water and left to soak over night so that sorters are not exposed to formalin vapour. Other fixation fluids that do not release formalin gas have been tested, such as formaldehyde depot chemicals (Dowicil 75 and Kohrsolin) used in clinics for sterilization purposes. The effects of these fluids on dry weight and ash-free dry weight are marked and the effects on long-term storage are unclear, so that no unequivocal recommendation can be given (Brey, 1986).

StainingTo facilitate sorting and to increase sorting accuracy, especially for small animals, staining the sample with, e.g., Rose Bengal, is recommended. However, in some cases, staining may cause problems with species identification and the time gained during sorting will therefore be more than offset. Zoological museums will not accept stained material for taxonomic purposes. The following procedure has been shown to give good results:Wash the sample free from the preservation fluid by using a sieve with a mesh size smaller than 0.5 mm x 0.5 mm. Allow the sieve to stand in Rose Bengal stain (1 g dm-3 of tap water plus 5 g of phenol for adjustment to pH 4-5) for 20 minutes with the sample well covered. Wash the sample until the tap water is no longer coloured, As an alternative, Rose Bengal (4 g dm-3 of 40 % formaldehyde) may be added to the fixation fluid. Overstained specimens may be destained in alkaline (pH 9) fluids (Thiel, 1966).

Sieving of Fixed MaterialSamples may be sieved 'alive', as is the usual practice, or preserved. If they are preserved, it must be realized that the sorting characteristics are different from those for live fauna and result in apparently higher abundance and biomass figures. Intercalibrations of both procedures should be performed. In publications, it should always be stated whether the sieved material was fresh (alive) or fixed.

Sorting

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Sorting must be done using some magnification aid (magnification lamp, stereomicroscope). Any finer fraction (< 1 mm) should always be sorted under a stereomicroscope.When taxa occur in great numbers (e.g., Polydora, Phoronids, Capitellids), it may be advisable to split the samples to reduce the counting time. Different types of sample splitters can be used. Rare species should be counted from whole samples. The accuracy of the sample-splitting device should be adequately assessed. To reduce sorting time, a sorting aid (such as the one described by Pauly (1973) or a 'fluidized sand bath' (after P. Barnett, see Holme and McIntyre, 1984)) may be used, provided that its efficiency has been satisfactorily checked for the particular bottom material studied. The Ludox method (see Higgins and Thiel, 1988) has successfully been applied to meiobenthos work and may also prove useful for the extraction of soft-bodied macrofauna. In coarse sand, the following procedure may be recommended: the sediment is fixed and placed on a PVC trough 5 m long, 20 cm wide, and 20 cm high (an ordinary gutter of the same length may also be used). Water is poured over the sediment from one closed side and the extracted fauna caught on a sieve on the other (open) side (Vanosmael et al., 1982). If samples are sorted alive, care should be taken to avoid predation within the sample.

Biomass DeterminationThe following measures of biomass determination can be used: wet weight, dry weight, and/or ash-free dry weight, either from fresh or fixed material. Furthermore, energy content (J) and / or matter equivalents (C, N, P) may be determined, using fresh material only. Fresh wet weight is to be preferred to formalin wet weight, but if the latter has to be used, weighing should not be done until at least three months after fixation (Brey, 1986).The wet weight is obtained by weighing after the external fluid has been removed on filter paper. The animals are left on the filter paper until no more distinct wet traces can be seen. Animals with shells are generally weighed with their shells; the water should be drained off bivalves before weighing. When shell-free weights are given, the shell weight should be included in the data list. Echinoids should be punctured to drain off the water before blotting on filter paper. As soon as the non-tissue water has been removed, the organisms are weighed with the accuracy required (for adult macrofauna: 0.1 mg). In case tube-building animals have to be weighed together with their tubes, appropriate correction factors should be established. The dry weight should be estimated after drying the fresh material at 60 oC, or by freeze drying, until constant weight is reached (at least 12-24 hours, depending on the thickness of the material; large bivalves may need up to 96 hours). Dry weights obtained by lyophilization (freeze drying) are slightly higher than those obtained by oven drying. For Mytilus, lyophilized tissues weighed 10.9 % more than oven-dried tissues (Gaffney and Diehl, 1986).The use of ash-free dry weight is recommended in routine programmes, because it is the most accurate measure of biomass (Rumohr et al., 1987; Duineveld and Witte, 1987). However, it destroys specimens, and the consequences of this should be carefully considered. Ash-free dry weight should be estimated after measuring dry weight. It is determined after incineration at 500 oC in an oven until weight constancy is reached (about 6 hours, depending on sample and object size). The temperature of the oven

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should be checked with a calibrated thermometer because there may be considerable temperature gradients (up to 50 oC) in a muffle furnace. Caution is advised to avoid exceeding a certain temperature (> 550 oC), at which a sudden loss of weight may occur owing to the formation of CaO from the skeletal material of many invertebrates (CaC03). This can reduce the weight of the mineral fraction by 44 %. Such decomposition occurs very abruptly and within a small temperature interval (Winberg, 1971). Before weighing, the samples must be kept in a desiccator while cooling down to room temperature after oven drying or removal from the muffle furnace.To estimate biomass from length or size measurements, conversion factors may also be used (Rumohr et al., 1987; Brey et al., 1988).

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A protocol for the study of meiofaunaUniversity of Crete

This protocol has been compiled from the following primary sources: Fleeger JW, Thistle D, Thiel H (1988) Sampling equipment. In: Higgins RP, Thiel H

(eds) Introduction to the study of meiofauna. Smithsonian Institution Press, Washington DC, London, p 115-125

Pfannkuche O, Thiel H (1988) Sample processing. In: Higgins RP, Thiel H (eds) Introduction to the study of meiofauna. Smithsonian Institution Press, Washington DC, London, p 134-145

Giere O (1993) Meiobenthology. The microscopic fauna in aquatic sediments. Springer-Verlag, Berlin, p 328

The Darwin Nematode Project website http://www.pml.ac.uk/nematode/index.htm

1. SAMPLING 1.1. Field equipment and materialsCorers Cylindrical, transparent, perspex tubes (proposed inner

diameter 4.4 cm) with smooth internal surface and beveled lower end to facilitate sediment penetration and core removal. The length should be at least 15 cm (Fig. 1a)

Rubber stoppers Appropriate diameter for tight-fitting to the coring tubes (Fig. 1)

Core plunger Same diameter with the corers for extruding the sediment core (Fig. 1b)

Corers basket Will ensure the upright position of the core samples and their carriage

Rubber mallet 45 um sieve Metal stainless steel or Nytex nylon gauzeSample containers ~ 500 ml volumeSample labels Indicate area, station, replicate and dateSyringe ~ 20 ml volume7% MgCl2 7.5 g MgCl2 6H2O dissolved in 100 ml distilled water. Used

as a narcotic agent10% buffered formalin Filtered seawater (through a 45 m sieve) should be(4% formaldehyde) used as dilutant to prevent contamination with planktonic

species. The formalin should be buffered with 200 g Borax per liter

Washbottles Used for distilled/filtered water, MgCl2

Spatulas Helpful for the slicing of the core

1.2 Sampling techniqueIn sediments, coring is the best quantitative sampling technique for meiobenthos, because when corers are used with care they collect a known area or volume of sediment with all depths equally represented and all animals present before sampling are captured.

Three general problems arise with core sampling:

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a) bow-wave-induced reduction in abundanceb) effects on population parameters due to the underlying distribution of the fauna (patchiness)c) sample distortion due to core compaction

These problems are minimized by taking cores slowly and selecting an appropriate core size.

If at all possible, subtidal samples should be taken by SCUBA divers (Fleeger et al. 1988). Divers usually obtain superior samples because they are able to position the samplers with care and insert the corer slowly (McIntyre 1971). In addition, the presence of the investigator will often yield important insights about the ecology of the site or practical aspects of the sampling (Fleeger et al. 1988). However, if diving is not possible, a Craib type corer (Fig. 2a) may be used instead. Subsampling from a sample collected with a multiple SMBA type corer, grab or box corer is also an alternative (Fig 2b,c). In the above case, caution should be taken to avoid pseudoreplication by taking each subsample from different depoloyments.

Figure 2. Different samplers used on research vessels for meiobenthic studies. a) Craib corer, b) box corer, c) multiple corer.

1.2.1 Sampling procedure

Figure 1. a) Corers, rubber stoppers and b) core plunger for sampling meiobenthos.

a b

a bc

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1. The corer should be inserted slowly and smoothly into the sediment in a vertical position.

2. Facilitate penetration with the use of a rubber head mallet3. The depth of penetration into the sediment must exceed 10 cm3

4. When the desired depth is reached, close the corer’s upper end with a stopper5. Remove the corer slowly. Care should be taken to retain the core while removing

the corer from the sediment. If needed, remove the sediment around the corer by digging for facilitating core removal.

6. Close the lower end of the corer with a stopper and place it in the basketCaution must be exercised to ensure that the corer will not be turned up-side down!

7. Use a separate, appropriately labeled container for each replicate sample 8. With the use of a syringe transfer the overlying water of each core in the sample’s

container9. Use the core plunger to extrude the core. If the surface layer is poorly

consolidated and cannot be extruded without loss (i.e. soft mud), the core can be allowed to slip down the corer by loosening the top stopper

10. Store in the container the top 10 cm of the core. Markings at appropriate intervals on the corer help identify the desirable thickness of the sediment

11. Add 7% MgCl2 until the sample is fully covered. Stir gently and allow 10 minutes to react

12. Fix the sample with 10% buffered formalin (for the dilution take into consideration the total volume of sediment)

13. Invert the container several times to mix the fixative and sediment.

2. SAMPLE PROCESSING 2.1. Equipment and materialsStereomicroscope45 µm and 0.5 mm sievesLudox TM solution Specific gravity ~1.15 Washbottles For filtered/distilled water and Ludox solution Containers or beakers The volume should be approximately five times the volume

of the sample Hydrometer A wine and beer maker's hydrometer is adequate (specific

gravity 1.10-1.16 approximately)Small containers 100 ml cup with screw cap can be used Rose Bengal solution 1 g Rose Bengal in 1 lt of 10% formalin10% buffered formalin(4% formaldehyde)

2.2 Meiofauna extractionMany of the following techniques involve washing and concentrating meiofauna on sieves with freshwater. Prior to processing samples it is important to check that the laboratory freshwater supply does not contain meiofauna. To do this run tapwater through

3 Required sample’s depth is 10 cm, but due to possible sediment loss that might occur while removing the corer from the sediment, the corer should be pushed deeper in the sediment

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a 45 µm sieve for 5-10 minutes and check the contents of the sieve under a stereomicroscope. If meiofauna is present attach plankton net of maximum 45 µm mesh size at the freshwater tap. It is also highly recommended to attach a flexible tube to the freshwater tap, as it greatly increases one's control over the direction and strength of the flow. Meiofauna are small, so do not use a strong jet of water, and splashing must be avoided.

2.2.1 Sample sievingSediment samples contain formalin and several sediment components such as silt, clay, sand grains and organic detritus, in addition to the fauna; therefore, they have to be sieved to extract a certain size fraction and remove the fine sediment and much of the formalin before organism concentration and processing. Sieving must be performed very carefully as rough handling might damage the animals.

2.2.2 Ludox flotation Most samples consist of a large amount of residue even after sieving and animals have to be extracted by density gradient separation. A number of flotation techniques have been used but a modification of the method proposed by de Jonge & Bouwman (1977) has been found to be simple and effective.The objective of flotation extraction is to suspend the fauna in a fluid which has a specific gravity very close to that of the animals themselves so that the animals are neutrally buoyant and remain in suspension but sediment components are negatively buoyant and slowly sink. A variety of dense fluids have been used to extract fauna from sediments in the past but nowadays most laboratories use Ludox, a colloidal silica solution, primarily developed for use in iron foundries, with properties which make it ideal for the extraction of meiofauna. It is available in a range of grades, but Ludox TM is most widely used.As supplied, Ludox TM has a specific gravity of 1.3 to 1.4. The specific gravity needed to extract meiofauna is approximately 1.15, so the stock solution must be diluted. Adding two parts fresh Ludox to three parts freshwater will give a solution of approximately the correct density, but the density should be measured with a hydrometer. Do not use seawater directly with Ludox, as this can cause the suspended silica to precipitate, rendering the sample useless.

2.2.3 Extraction procedure1. Wash the sample through a 0.5 mm sieve nested on top of a 45 µm sieve2. Large quantities of sediment, especially silt or clays, should be split into small

portions and wash without forcing through the sieve3. Extreme care is necessary not to clog the smaller mesh as it is not in direct view

Puddling may help if the sieve becomes clogged4. Continue washing until the water passing through the sieve is relatively clear5. Carefully wash the extracted portion of the sediment to one side of the sieve, then

wash it into a container using Ludox TM6. Add at least 4 times the sample volume of Ludox solution7. Stir vigorously to distribute the sample evenly throughout the volume and leave to

settle for 60 minutes. Some fine clay particles may remain in the sample even after

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washing and these will still remain in suspension even after several hours. They should be ignored

8. Carefully pour the supernatant through a 45 µm sieve into a jug9. Return the Ludox to the sample and repeat the flotation process (step 7-8) 5 times10. After each flotation wash the extracted fauna thoroughly with fresh water and

keep it to a cup labeled appropriately11. To facilitate animal detection, add in the entire sample a few drops of Rose

Bengal solution (24 hours is required to insure sufficient staining of meiofauna)12. Preserve the sample by adding formalin (the final concentration should be 10%).

3. PROCESSING MEIOFAUNA3.1 Equipment and materialsMagnetic stirrer100 ml beakerSmall magnetic stirring bar10-50 ml syringeStereomicroscope Magnification range 10x – 50xMicroscopeCounting trays Petri dishes marked with parallel lines / Bogorov’s tray45 µm sieveNeedlesMicroscope slides Coverslips WashbottlesGlycerol

3.2 Subsampling Typically, large sediment samples are required for quantitative work because of the small scale patchy distribution of meiofauna. Therefore, sediment samples contain large numbers of meiofaunal organisms, and it would be impossible to count and identify all of them. This is why we routinely identify a proportion, extracted at random from the whole sample, which we refer to as a subsample. As a general rule, a subsample which contains at least 500 specimens is adequate for standard community analyses. It is helpful to keep the subsample size (defined as a fraction of the whole sample) constant within a particular study.The Askö splitter (Elmgren 1973) was designed specifically for meiobenthic studies. However, a simpler subsampling method which performs sufficiently uses a magnetic stirrer and a small beaker in which freshwater is added to the sample up to a known volume.

3.3 Sample examination

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Figure 3. Counting trays for the examination of meiofauna under stereomicroscope. a) petri dish with grids, b) modified Bogorov’s tray.

Meiofauna should be examined and counted under a stereomicroscope at a minimum of 25x magnification. The smallest meiofauna (tardigrades, loriciferans, juvenile stages) are often only seen using 50x magnification. Specially designed counting trays, Bogorov’s tray or Petri dishes marked with squares on the bottom facilitate accurate scanning. Identification to higher taxonomic levels might sometimes require the specimen to be removed and examined under a microscope. In such a case, the extraction of the tiny specimen from the sample may be done with a fine needle, fine hooked needle, or with Irwin loop.

3.4 Procedure1. Wash the sample on a 45 µm sieve with freshwater2. Wash the sample from the sieve into a counter tray and check the size (in terms of

number of meiofauna organisms present) of the sample3. If the sample contains a large number of organisms (> 500) follow steps 4 – 7 for

subsampling; otherwise, go directly to step 84. Wash the sample into the beaker and fill it up with tap water to 100 ml volume5. Add a small magnetic stirring bar and place the beaker on the magnetic stirrer6. The key to efficient subsampling is stirring the sample gently using an average

stirring speed (~ 800rpm). It is also strongly recommended to shake the beaker by hand in a circular way when starting the stirring

7. After 10 seconds draw out with the syringe a certain volume4 of the sample avoiding the eddy

8. Transfer the sample/subsample into a counting tray (Fig. 3)9. Sort and count the sample/subsample into major meiofaunal groups5 under a

stereomicroscope at a minimum of 25x magnification. For very small individuals use higher magnification

4 The volume of the subsample will depend on the initial size of the sample

5 Nematodes, copepods (if possible count separately the different stages), annelids, isopods, amphipods, mollusks, turbellarians

ab

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10. For identification of tiny specimens, if needed, pick them out using a fine needle and place them separately in a drop of glycerin/water mixture on a microscope slide. Cover the specimen with a coverslip and examine it under a microscope.

Literaturede Jonge VN, Bouwman LA (1977) A simple density separation technique for

quantitative isolation of meiobenthos using the colloidal silica Ludox-TM. Marine Biology 42:143-148

Elmgren R (1973) Methods of sampling sublittoral soft bottom meiofauna. Oikos Supplement 15: 112-120

Fleeger JW, Thistle D, Thiel H (1988) Sampling equipment. In: Higgins RP, Thiel H (eds) Introduction to the study of meiofauna. Smithsonian Institution Press, Washington DC, London, p 115-125

Pfannkuche O, Thiel H (1988) Sample processing. In: Higgins RP, Thiel H (eds) Introduction to the study of meiofauna. Smithsonian Institution Press, Washington DC, London, p 134-145

Giere O (1993) Meiobenthology. The microscopic fauna in aquatic sediments. Springer-Verlag, Berlin

McIntyre AD (1971) Deficiency of gravity corers for sampling meiobenthos and sediments. Nature 231:260

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Protocols for Acoustic and Visual Surveys of the Seabed

Acoustic ground discrimination systems (seabed classification systems)Acoustic ground discrimination systems allow for mapping bathymetry, sediment type and some types of habitats (vegetation, mussel beds) in a range of a few kilometres around the planned location of aquaculture facility. For a proper sampling design, the desired spatial resolution of the map should be defined in advance, e.g. an area of a certain seabed type and a maximum diameter of 10 m should be identified with a probability of at least 90 % (Fig. 1). This can be achieved by choosing the distance between tracks scanned by the echosounder appropriately. Depending on the system in use a more or less extensive ground truthing (diver observations, photo, video or grab samples) is necessary to link sea bed types with habitat types.

Fig.1. Hypothetical track (red) of an echosounder. The sampling grid must ensure that discriminable seabed types of a certain size are detected with a reasonable high probability (e.g. 90 %).

Standard Operational Procedures vary with the technique, hard- and software in use, thus a thorough training of staff is necessary. Systems based on multibeam technology or sidescan sonar allow for a faster and more accurate mapping but are more expensive and thus less widely available. The vendors of acoustic ground discrimination systems offer manuals and training courses. Comprehensive guidelines on seabed mapping using acoustic ground discrimination interpreted with ground truthing have been published in the Marine Monitoring Handbook (Procedural Guideline No. 1-3, pages 183-197, No. 1-4, pages 199-209). The following protocol for using RoxAnnTM is based on a protocol written by Chris Cromey for the mobile equipment available at DML for use on an open vessel. Since the exact design of the equipment will vary for other installations, the protocol needs to be adjusted accordingly.

Objectives: To obtain bathymetry and sediment data for the site.The system can also be used to obtain accurate positional information for cage groups.The bathymetry information can be used in modelling and the sediment type data in general site assessment.

Deliverables: Bathymetry and sediment type data around fish farm area.

Personnel: 2

Time to complete task: 6 to 8 hours

Equipment:vessel

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minimum length ca. 5 m, exclusively available for the duration of the RoxAnn survey

supply of appropriate fuel hand held equipment for ship to ship communication lifejackets and normal safety equipment,

RoxAnn ground master system with cables (check!) transducer DGPS & GPS antennas & bracket Windows computer with dual comm ports and power supply waterproof computer box containing all comm. leads and video connections slave LCD screen AC/DC power supply and cables waterproof mouse sprayproof 240v distribution board with voltage filter adequate power generator removable data storage media transducer/antenna mounting bracket including packing transducer mounting pole antenna mounting pole tools! (all necessary screwdrivers, spanners etc.)

Setting up transducer and antenna bracket

Fit the vessel mounting bracket to side of vessel prior to adding transducer or mounting antenna. Adjust the spacing blocks as necessary to ensure a secure fit. Ensure that the assembly is as level as possible, thus ensuring that the transducer and antenna are as vertical as possible. Tighten the clamps, making sure that the outer struts are butted firmly against the retaining pegs, and all four of the struts are tightly clamped onto the vessel.

Secure the fore and aft ropes to the bracket on the transducer-mounting pole. Secure the transducer to the transducer-mounting pole. Best results are achieved from the system when the transducer is mounted as deep as possible in the water, taking account of minimum water depths that will be encountered during the survey. It is imperative that the transducer head does not become scratched or damaged in any way. Secure the transducer and mounting pole assembly to the vessel-mounting bracket. The direction of travel of the transducer head is important and the transducer should be rotated such that the point of the transducer head is facing aft, 180 to the direction of travel.

Attach the receiver beacons to the antenna mast. It must be ensured that the beacons have an unobstructed line-of-sight view of the sky, i.e., the beacons must be at least the same height as any other structure on the vessel. In a small boat it is sufficient to situate the beacons above standing head height.

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Setting up GroundMaster system, computer, and monitor

GeneralThe GroundMaster, the computer and the slave screen are all powered by 240v (13A), and therefore the plugs must be kept dry at all times. The plugs are configured to fit into a sprayproof distribution board that has an inline RCD and a mains filter to smooth the electricity supply to the equipment. The distribution board and plug are sprayproof only when the plugs are fully seated and connected. It is essential that only this distribution board be used for electricity supply to the system.

GroundMaster systemThe GroundMaster system should be kept dry at all times. Remove the lids from the two GroundMaster boxes and attach together at the hinges. Fix in the open position using the brackets (housed in the lower section). Do not allow the two halves to slam shut, get assistance if required. Place the frame over the GroundMaster and cover with the waterproof canopy.

Connect the two GroundMaster system boxes together using the cables. The cables are use specific and the plugs and sockets are not interchangeable. Only two cables share the same configuration of plug and socket (data cables – RS232 & Data out) and are interchangeable. Care must be taken when connecting the 24-pin SVGA cable, it is very fragile and the pins in the GroundMaster socket are very susceptible to damage.

Once all the cables are connected into the GroundMaster system (including transducer and DGPS beacon cables) there are 4 plugs leading from the system; RS232 & Data out; SVGA in and the 240v 13A power plug. The RS232 & Data out cables attach to the appropriately labelled ports on the waterproof computer box. The SVGA in connects to the video1 socket on the waterproof computer box. The 240v 3-pin plug connects to the specific distribution board.

ComputerEnsure both dongles (white for charts software and silver for RoxMap scientific software) are located in the parallel port of computer. Ensure the SVGA cable and the external mouse cable (plugs located inside box) are correctly fitted into the back of the computer. Within the waterproof computer box, the RoxAnn (RS232) data is fed to comm3 (port on PCMCIA card nearest back of machine) and the DGPS data is fed to comm5 (port on PCMCIA card nearest front of machine). These should be connected and it will not be necessary to touch these connections.

Slave monitorThis should be set up so that the helm can clearly see the screen. The SVGA connection from the external slave monitor (permanent 10m extension) connects to the Video2 socket on the computer box.

GeneratorCheck fuel level.

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Check oil level.Operate according to user manual.It may be necessary to run the generator for ca. 5 minutes before putting load onto generator (i.e., plugging equipment in).

Equipment startupThe GroundMaster system, Fugro SeaStar DGPS and the slave monitor have no power switch, and therefore switch on as soon as external power is connected. It is therefore essential that all cable connections have been made and the power source has been allowed to stabilise (if required). The specific distribution board is still required whatever electricity supply is used.

The depth sounders cannot be switched on concurrently. If the sounder is on there is power being transmitted through the transducer. Do not switch the transducer on in air; the brass head must be submerged in water.

Power up the GroundMaster unit (plug in), switch on the desired echo sounder and the Koden GPS. Switch on computer and wait for it to start up. Note that the external monitors do not function correctly until the computer has fully booted and loaded Windows operating software. Once the GroundMaster system, echosounder, GPS have been started, the operation of the system is operated by the computer software.

Changes to GroundMaster system

With both echosounders switched off, the sounder frequency can be selected by turning knob between high (200kHz) and low (50kHz) frequency sonar. Once the correct frequency is selected the particular echosounder can be turned on.

The GroundMaster gain calibration settings must also be selected to match the sounder. The file C:\Roxmap\RoxAnn calibrations.doc indicates the GroundMaster gain settings, the echosounder settings, and the boxfile settings in the RoxMap software, specific to transducer frequency and bottom type being surveyed. It should be noted that the only setting of the echosounder that affects the GroundMaster system and RoxAnn signal is the range setting. Do not attempt to alter the GroundMaster gain settings or the echosounder range settings unless you have been specifically shown how to and requested to do so by an authorised person.

Computer use

The software used is RoxMap Scientific 32. Double click on the desktop icon to start the software. Accept the two disclaimers. The screen will show a chart. To select the survey area required, zoom to a big scale by clicking the left mouse button. Alternatively, to zoom in on a particular area click and hold down the right mouse button to draw a rectangular selection box over the required area.

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Click on the Displays menu and select the RoxAnn display (showing boxfile and E1 and E2 values) and the Depth display (depth and seabed type).

Once the survey has commenced and the data appears on screen (along with the survey tracks) that data will only be saved to disk if Saving is switched on. Prior to saving the data, confirm that the save rate is at the desired settings. To check (and alter) the save rate, go to Tracks\Options and select the time interval. Saving of the data can be switched on by two methods:

EitherBy depressing the third icon in from the top right of the RoxMap screen (showing a disk symbol)

OrBy using the bottom menu bars, selecting Tracks\Saving.

Using either method, the user is prompted to confirm if saving is to be switched on (if it is off) or off (if saving is already on). If the same buttons are selected again, this will turn saving off (user prompted to confirm).

Saving and exporting data

Once the days survey has been completed and the computer has stopped saving data, the data needs to be exported into a form that can be processed. To do this, whilst in the RoxMap Scientific 32 software, press ALT and the windows key, and from the Programs menu select RoxMap Scientific and then select Export Text (Database C__). A window will quickly show (Converting…) and then disappear. The database file containing the data in ASCII format (tab delimited) will have been saved to the C:\Roxmap\database.txt. It is important that this file is copied from that directory to a specific directory and labelled accordingly. If there are several days worth of day on the screen at once, all of this data will be saved to the file and therefore the specific data will need to be selected from that file. The output of the file does not contain column headings, but are listed below:

Roxcolour Latitude Longitude Depth (m)

E1 E2 Time (from satellite)

Date

To exit the RoxMap software, click Hide to remove all the menus at the bottom of the RoxMap window, and then click Q that appears in the bottom right-hand corner of the window.

Protocol for labelling and processing data files (“#” = character or number)

#####raw.txt = Original database.txt file from RoxMap exported as text file and renamed (use date as file name descriptor)

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#####ed1.xls = raw file imported into Excel. Date changed to actual date (format changed from dd-mm-100 (for Y2K) to dd/mm/yy) and time changed to real time (format hh:mm:ss, add 8 hrs to convert from RoxMap time GMT to GMT, add 7 hrs to convert from RoxMap time BST to GMT)

#####ed2.xls = Depth in ed1 file converted to total actual water depth (depth of transducer corrected for). Original (unedited) date and time columns to be deleted.

#####ed3.xls = ed2 file processed through RoxCon (Remove tidal height) to allow for tidal height variations during survey. Total actual water depth is corrected to chart datum depth for survey area.

#####ed4.xls = ed3 file run through RoxCon (Filter data) with Method 1 maximum set to shallowest depth (i.e., –2m) and minimum set to deepest depth (i.e., -40m). Additional spikes may be removed by Method 2 spikes using an equal setting (e.g., 8m) for the above and below values. Deleted data file produced by RoxCon to be added as worksheet within this Excel workbook file.

#####ed5.xls = Spikes in ed4 file visually edited. Deleted rows to be added to bottom of deleted data file worksheet from RoxCon.

#####ed6.xls = ed5 file tidied up and columns renamed. Only necessary data columns remain in file as shown below.

Roxcolour Latitude Longitude Chart datum depth (m)

E1 E2 Time (GMT)

Date

#####ed6.txt = ed6 Excel converted to text file for import into other software and backup transmission (if applicable). Final version of data saved as text.

A log of datafile manipulation should be kept (as text file) for each set of survey data edited. This would normally be on a daily basis, but if two or more days worth of raw data are combined prior to manipulation, the log should reflect how all the data was combined and then manipulated.

Include in top of file: Depth of transducer = -#.#m; Sounder frequency = ###kHzEdit 1Time corrected

Roxcolour Latitude Longitude Depth E1 E2 Time(RoxAnn)

Date Time (GMT) Date

Edit 2Depth corrected

Roxcolour Latitude Longitude Depth (m) Corrected (actual water)

depth (m)

E1 E2 Time (GMT)

Date

Edit 3Depth to CD (LAT)

Roxcolour Latitude Longitude Depth (m) LAT(m) E1 E2 Time (GMT)

Date Tid.Ht(m)

Edit 4Roxcon corrections

Roxcolour Latitude Longitude Depth (m) LAT(m) E1 E2 Time (GMT)

Date

Edit 5Visual corrections

Roxcolour Latitude Longitude Depth (m) LAT(m) E1 E2 Time (GMT)

Date

Edit 6 Roxcolour Latitude Longitude Chart E1 E2 Time (GMT) Date

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Datum depth (m)

Map datum:In the UK the survey positional information is in Latitude and Longitude (WGS84) which is then converted to National Grid co-ordinates which area linear scale.This assists in analysis and subsequent use of the data.It may be desirable to convert latitude and longitude positional data to a similar linear system if algorithms are available and can be adapted for such conversions of large data sets.

Video and Photo survey

Video can be used for interpreting and ground truthing data from an acoustic survey or as a primary survey technique for habitat mapping. If video is limited by turbidity good timing of the survey may improve the quality of a video survey considerably, e.g. in tidal areas water clearance is often best around slack tide. Good synchronization between video and GPS data is a prerequisite for mapping habitats.

Video cameras can be towed above the seafloor. In areas where obstacles (rocks, wrecks etc.) can be expected, the camera system should be protected by a frame (Figure 2). The frames can also be used for close-up inspections when placed on the seafloor. The resolution of still pictures from photo cameras is much better than the resolution of video footage. Thus, an additional photo camera facilitates the interpretation of videos considerably. The frame-camera should be towed at a constant distance above the ground. A weighted rope of known length within the view of the camera is a simple but very helpful way to achieve this goal. Alternatively, the video camera and accessory equipment (lamps, still photo) can be mounted on a sledge (Figure 3). Comprehensive guidelines for identifying biotopes using video techniques and in situ survey of sublittoral epibiota using towed sledge video and still photography have been published in the Marine Monitoring Handbook (Procedural Guideline No. 3-5, pages 241-251, and No. 3-14, pages 331-337). If a frame or sledge based camera system is not available and depth does not exceed ca. 30 m and only a small area needs to be surveyed for ground-truthing a hand-held video- or photo camera can be employed (see Marine Monitoring Handbook, Procedural Guideline No. 3-13, 327-330) or habitat classifications can be conducted by scientific divers who are familiar with habitats and species typical for the region (Marine Monitoring Handbook, Procedural Guideline No. 3-3, pages 233-239).

Transect Photography(this preliminary guideline for transect photography, which has been tested in MERAMED, will be elaborated if the method becomes a standard method in ECASA.)

For monitoring the impact of an existing mariculture facility a photographic documentation along a transect line has been shown to be a valuable tool. Necessary Equipment are standard diving equipment, a compass, a good visible transect line (e.g. yellow, minimum length 50 m, diameter ca. 3 mm) on a seawater-resistant reel, an

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underwater photo-camera (focal length should be not longer than 28 mm for 35 mm film) with an external strobe.

A lead weight (1 to 2 kg) is attached to the beginning of the line and the line marked with tape every 5 m, for distances longer than 50 m every 10 m and the distance written on the tape. Even if the number is not always readable on the photos, it is helpful for evaluation and the orientation of the divers. The led is placed at the point of maximum impact, e.g. in the centre under a fish cage. From there, the diver swims towards the desired direction, preferably with the current, thereby unrolling the transect line. While swimming back, the diver exposes a horizontal photo at each mark (with the mark visible in the picture, see Figure 4) for qualitative overview pictures. If quantitative pictures are desired (e.g. for counting the number of feed pellet) vertical pictures should be taken with a ruler within the picture or a photo camera with a frame should be used.

Figure 2. Video Camera within a closed (left) or open (right) frame. The frame protects the fragile equipment from damage. Camera and lamp are mounted on a pan/tilt head for adjusting direction of view. A photo camera is added for high resolution still pictures (pictures from Rumohr 1995).

Figure 3. Dismountable aluminium video sledge (modified Aberdeen type, pictures from Rumohr 1999).

Figure 4. Images along a transect line to document the near-range impact of a mariculture facility.

Literature

Davies J, Baxter J, Bradley M, Connor D, Khan J, Murray E, Sanderson W, Turnbull C and Vincent M (2001) (eds.) Marine Monitoring Handbook. http://www.jncc.gov.uk/page-2430, ISBN 1 86107 5243.

Rumohr H (1995) Monitoring the marine environment. Sci.Mar. 59 (Suppl.1) 129-138.

Smith C, Rumohr H (2003) Imaging Methods. In: AD McIntyre, A Eleftheriou (eds) Methods for Study of Marine Benthos. Blackwell.

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Measurement of Particle Flux from Fish Cages and Consumption of Feed Pellets by Wild Fish (Sediment Trap Approach)

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Title of your method/discipline

Measurement of Particle Flux from Fish Cages and Consumption of Feed Pellets by Wild Fish (Sediment Trap Approach)

Institute responsible IfMContact person (with email address)

Helmut Thetmeyeremail address: hthetmeyer@ifm.uni-kiel.de

DescriptionKeywords summarising analytical parameters and utility of method

Aquaculture, net-cages, environmental impact, sediment traps, uneaten feed pellets, wild fish, fish faeces, particle flux.

Expected results Proportion of uneaten feed released to the environment, proportion of uneaten feed consumed by wild fish, distribution of feed and faeces under net cages (e.g. g solids m-2 bed day-1).

Reliability for assessing aquaculture effects

Sediment traps have been successfully used to assess the amount of uneaten feed pellets and faeces affecting the sea floor below fish net cages (Dougall and Black 1999). The procedure described here is an extension of existing methods that allow taking an ecological peculiarity at Mediterranean fish cage farms, the aggregation of wild fish, into account.

Nature of samples/analyses e.g.. visual, in situ, modelled, chemical/biological data etc.

The sediment traps are used to sample solid, organic material (mainly feed pellets and faeces) in-situ to assess the chemical and ecological impact of fish farms. The analysis of trap contents is gravimetrical and chemical (CHN). The proportion of wasted feed consumed by wild fish is calculated by difference between two sets of traps.

Quantitative, qualitative or semi-qualitative?

Quantitative data: sedimentation rate (dry weight, organic carbon, and nitrogen), proportion of feed and faeces in particle flux, amount of feed wasted, proportion of wasted feed fed by wild fish.

Validation Correct placement of traps should be validated by modelling the distribution of feed pellets and faeces. As model input topographical information (at least mean depth), contemporary measurements of current speeds, and experimental data on settling velocities of feed and faeces are needed. Further validation can be achieved by comparisons with independent methods, e.g. sonar, visual counts, or tracer techniques. Testing the suitability of TiO2 marked food pellets to assess consumption by wild fish is part of the MERAMED project.

Method description, instrumentation and reference to SOP

SOPs for assessing the particle flux below net cages have been designed mainly with salmon culture in mind. These procedures neglect consumption of wasted feed by wild fish which has been regarded to be of minor importance in northern Europe. However, there is evidence that wild fish play an important role

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in the flux of solid wastes at Mediterranean fish cage farms.The method is based on the collection of solid material below net cages using traps (containers) with a known mouth area. The trap design should allow a horizontal orientation of the collecting surface and minimise resuspension of settled material. In order to take the consumption of lost food pellets by wild fish into account two sets of traps are deployed, one set directly below the bottom of the cage (upper traps), the other on the sea floor (ground traps). The estimated amount of pellets consumed by wild fish is calculated by difference in contents of upper and ground traps.

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Sampling device (strategy), instrumentation and sampling area (and no. replicates etc)

The traps are simple PVC cylinders mounted on a square base plate. To avoid resuspension it is important that the ratio between lenght and diameter is at least 5:1. (e.g. diameter 15 cm, height 75 cm). Each upper trap is attached to the cage with three thin nylon lines (“crowfoot”). The orientation of the trap is stabilised by a lead weight (1 to 2 kg) below the base plate. The ground traps are stabilised by 2 lead weights (1 kg each) and/or two rods (e.g. 55 cm length, 5 mm diameter) (Figure 1).6 upper traps and 6 ground traps are deployed by a SCUBA diver. The distance between traps should be 2 to 4 m. The traps are placed along a line parallel to the current direction. For the placement of ground traps, the drift of pellets should be estimated. A sketch with the positions of the traps relative to the cage has to be drawn. During installation the traps should be covered with a lid, and the lid shouldn’t be removed before raised sediments have disappeared. After 24 hours the traps are sealed with a cap and brought to the surface. Standard procedures are used for filtering (coarse glass fibre filters, 10 to 15 cm diameter, e.g. Whatman GF/A, Cat No 1820 150), drying at 60 °C, and weighing (accuracy 0.01 g).To determine the ratio between collected feed and faeces a carbon-nitrogen analysis is conducted. Therefore, the filters are thoroughly homogenised (e.g. powdering with mixer “IKA A11 basic” and fine grinding with “IKH Ultra Turrax”) and dried at 60 °C. Subsamples are transferred into silver capsules, inorganic carbonate removed through dissolution in HCl, dried, and analysed in a CHN analyser. The ratio between feed and faeces is calculated by comparison with C/N data of pure feed and faeces samples.

Special requirements diving equipment, sediment traps, drying oven, balance, instruments for homogenisation, filter equipment, CHN analyser, current meters.

Extra help needed Support for dive team, especially supply with compressed air.Current speed should be measured contemporary, so that the proper placement of traps can be validated. A meteorological station to measure wind speed and direction may be helpful in order to correlate current and wind data.

Conditions that limit sampling

– High waves, strong wind or currents, heat, boat traffic, deep water.

Deployment and removal of traps should not be conducted when sedimentation rate is very high (i.e. during feeding times and 30 minutes thereafter)

Traps need to be left out for 24 hours. Undisturbed conditions are required. Sampling activities

which may affect the feeding behaviour of wild and cultured fish should be avoided.

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Sampling activities which cause a suspension of sediments should be avoided.

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Approximately how much time do you need per site

2 x 2 hours for deployment and removal of traps, 3 hours for filtration and weighing, 5 hours for CHN analysis.

Field workNo. sampling stations 1 to 2 cages per farmBasis for design of sampling network/ transect

Discontinuous transects or grids

Info required in the field log (please add/delete as required, this is just a starting point, and orientated to benthic sampling)

– name of diver and leader of operation– trap identification code– geographical co-ordinates for each sampling station;– map/sketch with trap positions drawn in– bathymetric map, at least depth under cage– depth of traps (measured by divers at mouth of traps)– cage size– stocking of cage (species, size, stocking)– specifications of food pellets (type, size, settling speed)– specifications of feeding (feeding times, duration of feedings,

rations) – date and time of deployment and removal of traps current strength and direction disturbances (e.g. boat traffic) need to be noticed diver observations (e.g. abundance and feeding behaviour of

wild fish)

Initial description of sample(as above, this serves as an example, and refers to benthic sampling – please alter to your needs)

– if possible, counts of pellets in traps– problems, remarks– diver observations

Literature references Dougall NM; Black KD (1999) Determining sediment properties around a marine cage farm using acoustic ground discrimination: RoxAnn™. Aquacult. Res. 30:451-458.

Practical issuesDisinfection requirements, SOPs and legal requirements you follow in your country

Standard disinfection procedure for diving equipment and boatsRegulations for driving motor boatsSecurity regulations on motor boatsRegulations for scientific diving (Greek or German)Diving permissions from Greek authorities

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Practical information (for use in cost-benefit)

– 500 to 1000 EUR for traps– 250 to 2000 EUR for filtration unit– 700 EUR for oven– 500 EUR for balance– auxiliary services: boat, dive team diving equipment,

compressed air, fuel, freshwater to rinse equipment, current meter, meteorological station (wind speed and direction), equipment for homogenisation, CHN analyser

– minimum 2 x 2 hours for deployment and removal of traps, 2 hours for preparation of diving gear and boat, 3 hours for filtration and weighing, 5 hours for CHN analysis

Any other comments Patchy distribution or low quantity of lost pellets may negatively affect accuracy; pellet counts may be biased if pellets degrade rapidly. Fouling on nets and dead fish on the bottom of the net may be a source of particles. If possible, experiments cages with clean nets should be chosen for experiments. Particles found in the traps which are obviously different from feed and faeces (e.g. algae, shells, animals, scales and bones) should be removed from the filters and/or their presence noted.

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Measurement of Particle Flux from Fish Cages with Sediment Traps without the Need of SCUBA Diving

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Title of your method/discipline

Measurement of Particle Flux from Fish Cages with Sediment Traps without the Need of SCUBA Diving.

Institute responsible IfMContact person (with email address)

Helmut Thetmeyeremail address: hthetmeyer@ifm.uni-kiel.de

DescriptionKeywords summarising analytical parameters and utility of method

aquaculture, net-cages, environmental impact, sediment traps, uneaten feed pellets, fish faeces, particle flux

Expected results Proportion of uneaten feed released to the environment, distribution of feed and faeces under net cages (e.g. g solids m-2 bed day-1).

Reliability for assessing aquaculture effects

Sediment traps have been successfully used to assess the amount of uneaten feed pellets and faeces affecting the sea floor below fish net cages (Dougall and Black 1999). The procedure described here allows for the deployment of traps below fish cages without divers.

Nature of samples/analyses e.g.. visual, in situ, modelled, chemical/biological data etc.

The sediment traps are used to sample solid, organic material (mainly feed pellets and faeces) in-situ to assess the chemical and ecological impact of fish farms. The analysis of trap contents is visual, gravimetrical and chemical (CHN analysis).

Quantitative, qualitative or semi-qualitative?

Quantitative data: sedimentation rate (dry weight, organic carbon, and nitrogen), proportion of feed and faeces in particle flux, amount of feed wasted.

Validation Validated by modelling the distribution of feed pellets and faeces. Contemporary measurements of current speeds and experimental data on settling velocities of feed and faeces are needed as input of the model. Further validation can be achieved by comparisons with independent methods, e.g. sonar, visual counts, or tracer techniques.

Method description, instrumentation and reference to SOP

The method is based on the collection of solid material below net cages using traps (containers) with a known sampling area (Dougall and Black 1999). The trap design allows for a horizontal orientation of the collecting surface and minimise resuspension of settled material. The traps can be deployed without assistance by SCUBA divers.

Sampling device The traps are simple PVC cylinders mounted on a base plate. To

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(strategy), instrumentation and sampling area (and no. replicates etc)

avoid resuspension it is important that the ratio between lenght and diameter is at least 5:1. (e.g. diameter 15 cm, height 75 cm). For vertical orientation of the traps in a current lead weights (1 to 2 kg) may be attached to crowfoots at the bottom of the traps. A rope is prepared as shown in the figure below which is marked according to dimensions of the cage and is lightly weighted at g and h. Prior to attaching the sediment traps, the rope needs to be positioned as shown in the figure. This can be undertaken by lowering the whole rope at position x but retaining rope ends on the surface, so that marks a and b are well underwater and g, h and m are close to or on the sea bed. The rope end closest to mark b can then be walked around the cage to position y. The weights at g and h close to the sea bed prevent snagging of the rope with the cage bottom. Once at position y, the rope ends can be hauled up both sides so that marks a and b are showing as in the figure. To attach the traps, the rope is then hauled out at y so that mark g passes underneath the cage and arrives at position y. About 12 sediment traps should then be attached at equal spacing between marks g and h as the rope is lowered at y and hauled in at x. Care must be taken not to haul in at x too tightly as to snag the cage bottom. Once marks a and b show, deployment is complete. After 24 hours the traps are brought to the surface.

A sketch with the positions of the traps relative to the cage has to be drawn. Standard procedures are used for filtering (coarse glass fibre filters, 10 to 15 cm diameter, e.g. Whatman GF/A, Cat No 1820 150), drying at 60 °C, and weighing (accuracy 0.01 g).To determine the ratio between collected feed and faeces a carbon-nitrogen analysis is conducted. Therefore, the filters are thoroughly homogenised (e.g. powdering with mixer “IKA A11 basic” and fine grinding with “IKH Ultra Turrax”) and dried at 60 °C. Subsamples are transferred into silver capsules, inorganic carbonate removed through dissolution in HCl, dried, and analysed in a CHN analyser. The ratio between feed and faeces is calculated by comparison with C/N data of pure feed and faeces samples.

Special requirements a long rope or series of ropes of total approximate length (3 times cage depth + 3 times cage length), sediment traps, drying oven, balance, instruments for homogenisation, filter equipment, CHN analyser

Extra help needed Current speed should be measured contemporary. A boat is needed to reach the cages.

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Conditions that limit sampling

– High waves, strong wind or currents. Deployment and removal of traps should not be conducted

when sedimentation rate is very high (i.e. during feeding times and 30 minutes thereafter)

Traps need to be left out for 24 hours. Undisturbed conditions are required. Sampling activities

which may affect the feeding behaviour of the cultured fish should be avoided.

Sampling activities which cause a suspension of sediments should be avoided.

Approximately how much time do you need per site

2 x 1 hour for deployment and removal of traps, 3 hours for filtration and weighing, 5 hours for CHN analysis.

Field workNo. sampling stations 1 to 2 cages per farmBasis for design of sampling network/ transect

Discontinuous transects or grids

Info required in the field log (please add/delete as required, this is just a starting point, and orientated to benthic sampling)

– name of leader of operation– trap identification code– geographical co-ordinates for each sampling station;– map/sketch with trap positions drawn in– estimated depth of traps– depth under cage– cage size– stocking of cage (species, size, stocking)– specifications of food pellets (type, size, settling speed)– specifications of feeding (feeding times, duration of feedings,

rations) – date and time of deployment and removal of traps current strength and direction disturbances (e.g. boat traffic) need to be noticed

Initial description of sample(as above, this serves as an example, and refers to benthic sampling – please alter to your needs)

– if possible, counts of pellets in traps– problems, remarks

Literature references Dougall NM; Black KD (1999) Determining sediment

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properties around a marine cage farm using acoustic ground discrimination: RoxAnn™. Aquacult. Res. 30:451-458.

Practical issuesDisinfection requirements, SOPs and legal requirements you follow in your country

Standard disinfection proceduresRegulations for driving motor boatsSecurity regulations on motor boats

Practical information (for use in cost-benefit)

– 500 to 1000 EUR for traps– 250 to 2000 EUR for filtration unit– 700 EUR for oven– 500 EUR for balance– auxiliary services: boat, fuel, freshwater to rinse equipment,

current meter, meteorological station (wind speed and direction), equipment for homogenisation, CHN analyser

– minimum 2 x 1 hours for deployment and removal of traps, 3 hours for filtration and weighing, 5 hours for CHN analysis

Any other comments Patchy distribution or low quantity of lost pellets may negatively affect accuracy; pellet counts may be biased if pellets degrade rapidly.This method does not allow opening and closure of traps during deployment so if the nets are fouled there is potential for material from the nets entering traps. Care must also be taken to prevent bottom sediment entering traps. Alternative deployment methods include vertical deployment of traps at the side of cages, either inside or outside the nets although no traps can be deployed underneath the cage with this method. However, traps can be deployed inside the cages at the middle point by using a bar the length of l or using a central walkway. Particles found in the traps which are obviously different from feed and faeces (e.g. algae, shells, animals, scales and bones) should be removed from the filters and/or their presence noted.

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cage length, l

cage depth, d’

a

m

b

g h

rope

water column depth, d

x y

sea surface

sea bedFigure 1. Deployment of sediment traps without the use of SCUBA diving for a cage dimensions l and d’. The rope is marked at positions a, b, g, h and m according to these dimensions.

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