Book of Protocols
ECASA Project
1
Table of Contents
page
Table of Contents
Selection of sampling stations
Sampling Field Log
Hydrography
“Large-scale” description of the seafloor at the
study site, by visual & other means
Sediment sampling
Grab sampling
Benthos
Macrobenthos procedures
Meiofauna procedure
Phytobenthos procedure
Sediment chemistry
Sediment redox potential, Eh
Sediment grain size analysis (granulometry)
Sediment organic matter
Sediment organic carbon
Total nitrogen in sediment
Inorganic and organic phosphorus in sediment
Water column (water quality) sampling
Secchi disk transparency
Dissolved nutrients in seawater
Nitrate/Nitrite in seawater
Orthophosphate in seawater
Ammonium in seawater
Total N & total P in seawater
Particulate matter in seawater
Particulate organic carbon and nitrogen in seawater
Particulate organic phosphorus in seawater
Chlorophyll a determination
Bioassay studies to assess nutrient levels around
aquaculture
1
2
2
3
4
6
6
9
9
16
18
20
20
22
23
24
26
27
28
29
31
34
34
35
35
38
40
42
43
46
2
Selection of sampling stations
Station positions should reflect gradients of impact. The principle is that some rationale
must be used to maximise the chance that one of the stations is selected in the area of
maximum impact. This can be done by modelling (using an appropriate current record)
or by a pre-survey which could be done by sediment sampling for e.g. redox, by acoustic
methods or by video, etc.
We want to maximise the chance of detecting a gradient and we also would like to
consider the distance from the farm that effects of the farm can be detected. This will
probably take a minimum of 5 stations per site but could require several more.
A reference station should be at a similar depth and sediment type but not be influenced
by the farm. If this is not possible, an additional station added to the transect may reveal
background conditions by comparing the trend away from the farm i.e. if stations become
more similar with distance.
Sampling Field Log
One of the key elements necessary to ensure successful fieldwork is a good and reliable
system for data collection and handling. It is essential that careful records are kept of all
details related to the sampling and measurement of field data. A typical field log should
include:
– date of sampling;
– 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
– other comments (such as rejected samples, delays and any problems experienced and
the causes);
– quality of sample i.e. disturbed/not disturbed.
3
Hydrography
Source: MERAMED project field handbook
Hydrography is one of the key features used in site selection, as it indicates: a) flux of
good quality water via the finfish or shellfish gear (water quality, animal welfare); b)
spatial dispersal of effluents, which may spread the 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 before the
intensive WP5 site study as it will serve as the basis for pre-selecting sampling stations
(see below).
For microtidal sites (e.g. Mediterranean) current meter records should be as long as
possible; at the very least 15 days, but preferably months of data. For macrotidal sites
(e.g. France, Atlantic coast), 15 days may be sufficient to capture the tidal cycle.
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 minutes is recommended for long-term deployments (e.g. 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. Ushaped 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
4
- 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
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.
“Large-scale” description of the seafloor at the study site, by
visual & other means
Source: Davies et al. (2001)
Video or still photography of the seafloor, along a transect (from the fish/shellfish farm
toward the reference station) is extremely desirable as it will provide large-scale
information on macrophytobenthos, epibenthic macrofauna, seafloor status, etc. which
core or grab sampling cannot do. 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. 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 framecamera 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
5
this goal. Alternatively, the video camera and accessory equipment (lamps, still photo)
can be mounted on a sledge. 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. 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 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 legible on the photos, it is helpful for evaluation
and orientation of the divers. The weight 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) 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.
Acoustic ground discrimination systems (seabed classification 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 %. 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.
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. 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).
6
References
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.
Sediment sampling
Timing of sampling is crucial - sampling should be done when impact is likely to be
high.
Number of sediment samples. At each station, at least 4 grab replicates should be taken
for macrofauna, and triplicate cores for sediment chemistry. If an analysis is very
expensive or complicated to perform, duplicates may be sufficient.
Diver sampled cores. Sediment cores should be taken by divers where practical since
divers can apply quality control (i.e. discard bad samples and re-sample in situ) to the
sampling process. Core diameters should be 5cm for sediment chemistry and 10cm for
macrofauna. Core depth should be at least 12cm for sediment chemistry and at least 6cm
for macrofauna.
1. The corer should be inserted slowly and smoothly into the sediment in a vertical
position.
2. If necessary, facilitate penetration with the help of a rubber head mallet
3. The depth of penetration into the sediment must exceed 10 cm1
4. When the desired depth is reached, close the corer’s upper end with a stopper
5. 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 a carrying basket
Caution must be exercised to ensure that the cores are not be turned up-side down
during transport, as this will mix the enclosed sediments!
Grab sampling
Source: Stubbs et al. 1987
Where it is not possible to collect by diving, samples will generally be taken by van Veen
grab with a sampling area of 0.1 m2. Grab sampling is suboptimal for sediment
geochemistry or for meiofauna sampling because the stratified structure of the sediments
is usually compromised by the sediment collecting mechanism of the grabs. However,
1
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
7
because van Veen grabs are affordable and are easy to use from a small boat, these will
most likely be used by farmers and regulators monitoring sediments below fish farms.
Whereas box corers yield undisturbed sediments, they are not recommended in this
project because most farmers/regulators will not have access to box corers (expensive) or
to the large boats (with large crane/winch) needed to use these.
If possible, samples could be taken with a Craib corer as this yields undisturbed sediment
profiles.
Gravity corers and grabs tend to fluidize the samples and destroy the vertical
stratification of the sediments, thereby making chemical or meiobenthos data extracted
from them meaningless.
Benthic sampling using a Van Veen grab
A 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 by using a 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 procedures
Prior 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.
Criteria for acceptable grab samples
Upon 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.
8
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
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.
9
Sediment Description
The 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.
Precise position fixing during sampling is essential. The position and the depth should be
controlled and documented during station work.
References
Stubbs, H.H., D.W. Diehl, and G.P. Hershelman. 1987. A Van Veen grab sampling
method. So. Calif. Coastal Water Res. Proj., Long Beach, CA., Tech. Rep. No. 204.
Benthos
Benthos samples will consist of the entire grab (i.e. no a priori subsampling) or the entire
core. Macrofauna samples must be sieved sequentially at 1mm and 0.5mm and
macrofauna will be identified to lowest practical taxon.
Meiofauna will be taken from all possible sites by cores. If necessary, meiofauna may be
subsampled from van Veen grab, by using sub-sampling cores. Where possible,
meiofauna should be taken from cores collected by a Craib corer.
Macrobenthos Procedures
Assorted Sources
Separation of Fauna from the Sediment
The 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.
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
10
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 Relaxants
Relaxants 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 softbodied 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.
Recommended treatments for main marine groups:
Sponges: Fix and preserve in 5% formaldehyde. Calcareous sponges should be preserved
in 75% ethanol as formaldehyde can decalcify the specimens.
Hydroids: Relax in 8% MgCl2 (or 15% MgSO4 or Menthol crystals). Fix in 5%
formaldehyde for at least 24 hours; transfer to 75% ethanol for preservation
*Actinians: Allow to relax in seawater then narcotise by replacing slowly with either
8% MgCl2 or Soda water to 50% (or 10% MgSO4 plus 1 or 2 drops of formaldehyde
every 15 minutes)
Nemerteans: Relax in 8% MgCl2 or add Menthol crystals to water
11
*Polychaetes: Relax in 8% MgCl2 (or gradual addition of 70% ethanol, or 20% MgSO4,
or 0.15% propylene phenoxetol to water). Fix in 5% formaldehyde for 24 hours then
transfer to 1.5% propylene phenoxetol (this preserves colour, but if unavailable 75%
ethanol will do). Ideally, don't fix in ethanol and don't leave in formaldehyde.
*Priapulids, sipunculans, echiurans: Relax using menthol crystals with a few drops
of alcohol added after an hour (or put straight into 8% MgCl2)
Small Crustaceans: Relax in soda water (or add a few drops of 70% ethanol to water
or use 0.15% propylene phenoxetol).
*Opisthobranchs: Relax in 8% MgCl2, fix and preserve in 5% formaldehyde or transfer
to propylene phenoxetol after fixation.
Bryozoans: Calcified bryozoans fix and preserve in 75% ethanol, fleshy or membranous
ctenostomes fix in 5% formaldehyde for 24 hours then transfer to propylene phenoxetol
for preservation
Echinoderms: Fix in excess 75% alcohol, replace after a few days due to dilution from
body fluids. Do not preserve long term in formaldehyde as the acid can dissolve the
calcareous ossicles and plates, which are essential for identification, particularly of
holothurians.
*Ascidians: Relax using Menthol crystals (or immerse in 8% MgCl2). Fix in 5%
formaldehyde. They can be preserved in propylene phenoxetol, or left in formaldehyde.
*IMPORTANT to relax these groups BEFORE fixation if need to identify later
Sieving options
There 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
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 a 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
12
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
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).
13
Sieving Procedure
Sieving 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
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).
14
Staining
To 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.
Sieving of Fixed Material
Samples 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
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 Determination
The 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
15
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
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).
References
Brey, T. 1986. Estimation of annual P/B-ratio and production of marine benthic
invertebrates from length-frequency data. Ophelia Supplement, 4: 45-54.
Callaway, R., Robinson, L. & Simon P.R. 2003. Methods Manual, Managing Fisheries to
Conserve Groundfish and Benthic Invertebrate Species Diversity (MAFCONS
Project)
Duineveld, G.C.A., and Witte, H.J. 1987. Report on an intercalibration exercise on
methods for determining ash-free dry weight of macrozoobenthos. ICES CM
1987/L:39.
Gaffney, P.M., and Diehl, W.J. 1986. Growth, condition and specific dynamic action in
the mussel Mytilus edulis recovering from starvation. Marine Biology, 93: 401-409.
16
Gosner, KL 1971. Guide to identification of marine and estuarine invertebrates. John
Wiley, New York, New York, USA.
Higgins, R.P., and Thiel, H. 1988. Introduction to the study of meiofauna. Smithsonian
Institute Press, Washington, D.C. 470 pp.
Holme, N.A., and McIntyre, A. 1984. Methods for the study of marine benthos. IBP
Handbook, 16. Second edition. Oxford. 387 pp.
Pauly, D. 1973. Über ein Gerät zur Vorsortierungg von Benthosproben. Berichte der
Deutschen Wissenshaftlichen Kommission für Meeresforschung, 22 (4): 458-460.
Rumohr, H., Brey, T., and Ankar, S. 1987. A compilation of biometric conversion factors
for benthic invertebrates of the Baltic Sea. Baltic Marine Biologists, Publication No.
9. 56 pp.
Vanosmael, C., Willems, K.A., Claeys, D., Vincx, M., and Heip, C. 1982. Macrobenthos
of a sublittoral sandbank in the Southern Bight of the North Sea. Journal of the
Marine Biological Association of the UK, 62: 521-534.
Winberg, G.G. 1971. Methods for the estimation of the production of aquatic animals.
Academic Press, London, New York. 175 pp.
Meiofauna procedure
Source: University of Crete
Required
Corers
Rubber stoppers
Core plunger
Corers basket
Rubber mallet
Sample containers
Sample labels
Syringe
7% MgCl2
10% buffered formalin
(4% formaldehyde)
Washbottles
Spatulas
Cylindrical, transparent, plastic 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)
Appropriate diameter for tight-fitting to the coring tubes
(Fig. 1)
Same diameter with the corers for extruding the sediment
core (Fig. 1b)
Will ensure the upright position of the core samples
~ 500 ml volume
Indicate area, station, replicate and date
~ 20 ml volume
7.5 g MgCl2 6H2O dissolved in 100 ml distilled water.
Used as a narcotic agent
Filtered seawater (through a 45 um sieve) should be
used as dilutant to prevent contamination with planktonic
species. The formalin should be buffered with 200 g Borax
per liter
Used for distilled/filtered water, MgCl2
Helpful for slicing the cores
17
Sampling
In 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.
a
b
Figure 1. a) Corers, rubber
stoppers and b) core plunger
for sampling meiobenthos.
If 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 deployments.
a
b
c
Figure 2. Different samplers used on research vessels for meiobenthic studies. a) Craib
corer, b) box corer, c) multiple corer.
Processing cores
18
It is easiest to process the cores by working with another person. The bottom stopper is
removed and replaced by the core plunger, to avoid loss of the sample. Following this,
the upper stopper is removed and the overlying water is transferred, with a syringe, to a
sample jar (because meiofauna may have swum out of the sediments to the water). Next,
the sediment is pushed out of the corer and the top 10cm are transferred to the sample jar.
A 7% MgCl2 solution is added to the jar until the sample is fully covered. Stir gently and
allow 10 minutes to react. Next, the sample is fixed by adding buffered formalin to
achieve a final concentration of 10% (take into consideration the volume of sediment,
water and MgCl2). Invert the jar several times to mix the fixative and sediment and store
at room temperature until the samples may be processed further.
References
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. SpringerVerlag, Berlin
McIntyre AD (1971) Deficiency of gravity corers for sampling meiobenthos and
sediments. Nature 231:260
Phytobenthos procedure
Source: HIMOM
Background
Phytobenthos refers to the algae that may be found on/in the seafloor sediments.
Sediments are sampled and the algal pigments are quantified by spectrophotometer and
by HPLC. The sampling strategy is a key issue in these studies due to non-uniform
vertical and horizontal distribution of the phytobenthos. In muddy sediments,
phytobenthos is found mostly in the top 1.5 cm of sediment whereas in sandy sediments it
is possible to find phytobenthos as deep as 15 cm depth (Cartaxana et al., 2006).
Phytobenthos distribution may be very patchy and the sampling strategy should take this
into account.
No extraction procedure exists that will provide 100% extraction efficiency. Efficiency
varies with species and acts differently for the various pigments. The 90% acetone as
recommended in this protocol generally provides ca. 90% efficiency (Wiltshire et al.
2000). Moreover, acetone extraction is especially efficient for polar pigments, like
chlorophyll a, and thus can be used for the analytical methods described below.
Unfortunately, some extraction-resistant species such as cyanobacteria and Zostera exist
on the tidal flats, which require mechanical (e.g. grinding) and aggressive chemical (e.g.
toxic dimethylformamide and methanol) means to extract these pigments. However, due
19
to other problems associated with these extraction methods, they will not be described
here.
Required
- Freezer
- Freeze-Dryer
- balance
- 90% buffered acetone
- ultrasound bath
- centrifuge
- ultrasound bath
Sampling:
A small petri dish is used to take surface sediment samples at intertidal sites. Ideally,
the petri dish should have 20 ml volume and a height of 2 cm. The Petri dish is placed
on the sediment and collected by means of a plastic card that is slid under the plate to
break the contact with underlying sediment.
This method works very well with mud, however, for sampling sand, may want to
sample a deeper depth, based on previous (preliminary) experience regarding vertical
distribution of the phytobenthos. If sampling in a subtidal area, sediments may be
taken by divers, by Craib corer or by box corer (sediment surface must NOT be
disturbed) and then subsampled using the Petri dish method.
Lab work:
Extraction should always be carried out under dim light, keeping the samples and solvent
cold/on ice.
1. Sediment samples should be frozen as soon as possible and then freeze-dried
2. Weigh out an amount (about 15g) of freeze-dried sediment and transfer to 50ml
centrifuge tube
3. Add about 10% (w/w) quartz sand (MERCK- particle size 10-30µm) to the tube
4. Add 15ml acetone (10% saturated sodium carbonate : 90% pure acetone) to the
tube.
5. Place the tubes in an ultrasound bath containing -4ºC water for 90 minutes.
6. Place the sealed tube with sample in a freezer (-20ºC) to extract for 24 hours
(vortex after that).
7. At end of 24 h, vortex the tubes and centrifuge 10 min at 3000 rpm.
8. Take the extract for pigment analysis, using HPLC or spectrophotometer.
References
Bowles, N. D., Paerl, H. W. & Tucker, J. 1985. Effective solvents and extraction periods
employed in phytoplankton carotenoid and chlorophyll determinations. Can. J. Fish.
Aquat. Sci. 42: 1127-31
Cartaxana, P., Mendes, C.R., van Leeuwe, M.A & Brotas, V., 2006. Comparative study
on microphytobenthic pigments of muddy and sandy intertidal sediments of the Tagus
estuary. Est. Coast. Shelf Sci. 66: 225-230.
20
Wiltshire, K.H., M. Boersma, A. Molle & H. Buhtz, 2000. Extraction of pigments and
fatty acids from the green alga Scenedesmus obliquus (Chlorophyceae). Aquatic Ecol.
34: 119-126.
Wright, S. W., S. W. Jeffrey & R. F. C. Mantoura, 1997. Evaluation of methods and
solvents for pigment extraction. In: Jeffrey, S. W., Mantoura, R. F. C., and Wright, S.
W. [Eds], Phytoplankton pigments in oceanography: guidelines to modern methods.
UNESCO, Paris, pp. 261-82.
Sediment chemistry
At each station, triplicate cores will be taken for sediment chemistry. If an analysis is
very expensive or complicated to perform, duplicates may be sufficient.
Sediment chemistry will be conducted almost exclusively on sediment cores. The first
thing that will be measured in the sediments is the vertical profile of redox potential.
Cores (or sub-cores taken from grabs) will be sliced at 1 cm intervals from sediment
surface to 12cm depth for sediment geochemistry. If the redox discontinuity layer is
deeper than 12 cm, should slice down to the redox discontinuity depth.
Chemical variables will be measured from cores sampled by SCUBA divers or in subcores taken from grab samples. It is essential that grabs are of good quality for
chemical analysis, with as little disturbance as possible. If the samples have been
disturbed, best thing to do is toss the sample and sample again to obtain a good,
undisturbed sediment sample. If this is not possible, should note this and the reason
additional cores could not be taken and analyze only the top 2cm layer.
The sediment slices will be taken for granulometry, loss on ignition, total organic carbon,
total nitrogen, total phosphorus.
Sediment redox potential, Eh
Source: ICRAM
Organic enrichment of sediments usually leads to reduced conditions which equate to
“bad” sediment quality, wherein natural benthic communities undergo substantial
changes. The oxidation-reduction (redox) conditions in surface sediments depend on the
degree of organic enrichment and can be assessed by measuring the vertical redox
potential profile (expressed in mV) in the top 15 cm (Zobell, 1946). The redox state of
sediment is the result of the combined effect of biological and chemical processes of
reversible and/or irreversible nature and is therefore difficult to define. It has been
pointed out that the concentrations of the components reacting reversibly are, in most
cases, too small to get reliable results from redox measurements. Thus, what is really
measured is a mixed potential which is not useful for chemical equilibrium calculations
(Bågander, 1978). However, Eh profiles still provide useful information, since the
decrease in Eh with the depth is related to the decrease in the dissolved oxygen
21
concentration in the pore water. Negative Eh values are therefore associated with anoxic
conditions, in which the degradation of the organic matter is performed by anaerobic
bacteria. In marine sediments, these bacteria mainly use sulphate as the electron acceptor,
which is reduced to hydrogen sulphide (Porrello et al., 2005; Chamberlain, 2002;
Danovaro et al., 2004).
Redox potential is measured by profiling an electrode down a sediment core to as deep as
is necessary to detect the redox discontinuity layer (normally 10 or 15cm), i.e. the point at
which redox values change abruptly from highly negative values to either less negative,
or to positive values. Redox potential will be measured in duplicate cores at each station
and only in undisturbed cores.
Required:
Surface sediment in cores, at least 0-20 cm depth
Ruler (to measure sediment height)
Portable pH/Eh meter
Redox electrode, with shaft >15cm long, preferably as thin as possible, with Platinum
ring indicator
Double junction silver/silver chloride reference electrode
Redox electrode calibration:
The ZoBell solution is used as a reference; 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 redox potential should be measured by carefully pressing the redox electrode into the
intact sediment core (in the tube) as soon as possible after collection (the longer the
sediment sits in the collection tube, the more it changes due to biogeochemical processes
that directly affect redox values), at depth intervals of 1cm from the surface to 15cm
depth. Measuring redox by inserting electrodes into holes in the side of the cores leads to
artifacts (see Hionchey and Schaffner 2005) and is therefore not recommended.
Moreover, slicing the sediments to measure redox in these is not recommended as
exposure to the air will rapidly and drastically alter the redox potential values. It is
essential that the measurements be done using either a combination redox electrode or a
redox electrode that is connected to a reference electrode that is in contact with the
sediment porewater. The measurements will be recorded in mV when the meter readings
stabilize at each depth.
In order to clarify the effect of sampling method on the data, sediments should be
sampled (several replicates) at same site by both van Veen grab and by Craib corer or by
SCUBA diver sampled corers. Redox profiles should be run directly on the sediments in
the grab and subsequently, subsamples should be taken from the grab for redox profiles
and these should be compared to redox profiles performed on cores taken either by
SCUBA divers or by Craib corer.
References
22
Bågander, L.E. 1978. An evaluation of the use of redox measurements for characterizing
recent sediments. Estuarine and coastal Marine Science, 6: 127-134.
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.
Hinchey, E.K. and L. C. Schaffner. 2005. An evaluation of electrode insertion techniques
for measurement of sediment redox potential in estuarine sediments. Chemosphere
59:703-710.
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.
Sediment grain size analysis (granulometry)
Source: HIMOM
Introduction
The objective of grain size analysis is to determine the grain size characters of sediment
samples and subsequently classify samples based upon their constituent parts. Particle
size analysis is performed using laser diffraction techniques (<2mm fraction) and sieving
techniques (>2mm fraction). Sediments containing a mixture of both fractions are
analysed by both methods and the results integrated. The results from the particle size
analysis are used to generate size distribution plots, to calculate statistical operators
including mean and sorting, and to obtain %clay, %silt, %mud, %sand and %gravel in
order to classify samples into sediment types.
The laser diffraction technique is described in the HIMOM manual. The sieving
technique is executed as follows:
1. Put the sediment sample in an oven at 60-80.C to calculate total dry weight (it’s
advisable to take a sample of ca. 100g dry weight).
2. Remove the fraction <63um, by sieving the sample through a 63 um sieve under a
water flow.
3. Dry the remaining fraction in an oven at 60-80.C
4. Put this fraction in the analytical sieve shakers to separate the different fractions
(>1000um; 1000-500; 500-250; 250-125; 125-63um).
5. Weigh the residuals in each sieve, and express as a percentage of total weight,
including the weight of the fraction <63um, which is calculated by subtraction.
Go to HIMOM Book of Protocols, p. 45 for further details on granulometry.
23
Sediment organic matter
Ratio of refractory to labile organic matter using Loss on Ignition (LOI)
measurements
Source: Loh 2005 (SAMS)
Introduction
There are some data in the literature on LOI using this method, but generally little for fish
farms. It might be expected to work very well in fish farms owing to the high flux of
labile organic matter to sediments, and it should be possible to use this to evaluate
recovery status. This indicator is less complex and time-consuming than total organic
carbon (TOC) measurements, which are routinely carried out in EIA studies, yet correlate
very poorly with impact, as TOC includes both labile and refractory material.
Method
Loss on ignition is carried out in a temperature-monitored muffle furnace. Approximately
0.5 g of dried, ground and sieved (200um) sediment sample is weighed precisely into a
crucible. Crucibles with sediment are then ashed (250°C for 16 hours). When cooled, the
crucibles are reweighed. Sediments are then heated to 500°C (Kristensen and Andersen,
1987) for 16 hours (Sutherland, 1998). When cool, they are weighed again.
According to Kristensen (1990), the weight loss in the lower temperature range
(130-280°C) is due to evaporation (i.e. dehydrogenation of hydroxylated aliphatic
structures, decarboxylation of acid groups, and generation of low molecular weight
volatile compounds); oxidative degradative degradation of aliphatic carbohydrates;
random chain-scission of weak bonds; cross-linking and peroxide formation; formation of
compounds of less-ordered structure; cyclization; and formation of carbonaceous char.
Plant materials rich in carbohydrates are combusted at the lower temperatures. The
weight loss in the high temperature region (280-520°C) is due to oxidation of aromatic
groups (polyphenolic compounds such as lignin, humic substances and kerogens) and
char. The percentage weight reduction after both 250°C and 500°C are measures of the
sediment labile and refractory organic matter, respectively.
Mook and Hoskin (1982) found significant (p<0.05) weight losses between 200°C and
300°C; hence the mean 250°C is chosen to be used here. Kristensen and Andersen (1987)
found that most calcite was combusted between 500-800°C, hence the 500°C is used in
this study, as most organic carbon would be completely separated with this temperature.
Hirota and Szyper (1975) also separated organic and inorganic compounds by heating at
500°C.
Rp index was introduced by Kristensen (1990) to characterize the composition of various
biogenic organic materials at different decomposition stages. It is defined as, Rp =
PII/(PI+PII) where PI is the weight loss in the temperature range 0 °C and 250 °C and PII
is the weight loss in the temperature range 250 °C and 500 °C. Calculations of percentage
organic matter and Rp values are as follows:
24
Let Ws = original weight of sediment sample (approximately 0.5 g)
W0 = weight of crucible and sediment before combustion
W250= weight of crucible and sediment alter combustion at 250°C
W500= weight of crucible and sediment after combustion at 500°C
PI = weight loss in temperature range 0°C and 250°C = WO - W250
PII = weight loss in temperature range 250°C and 500°C = W250 - W500
and, Rp= PII/(PI+PII)
% labile OM = (PI/Ws) x 100%
% refractory OM = (PII/Ws) x 100%
% total OM =
W0-W500
Ws X 100%
or, % total OM = % labile OM + % refractory OM
References
Hirota, J. & J. P. Szyper, 1975. Separation of total particulate carbon into inorganic and
organic components. Limnol. Oceanogr. 20: 896–900.
Kristensen, E. 1990. Characterization of biogenic organic matter by stepwise
thermogravimetry (STG). Biogeochemistry 9:135–159
Kristensen, E. and F.Ø. Andersen. 1987. Determination of organic carbon in marine
sediments: a comparison of two CHN-analyzer methods. J. Exp. Mar. Biol. Ecol. 109:
15-23.
Loh, P.S., 2005. An assessment of the contribution of terrestrial organic matter to total
organic matter in sediments in Scottish sea lochs. PhD thesis, UHI Millenium
Institute, 350 pp.
Mook, DH and Hoskin CM. 1982. Organic determinations by ignition: caution advised.
Est. Coast. Shelf Sci. 15: 697-699.
Sutherland, R. A., 1998. Loss-on-ignition estimates of organic matter and relationships to
organic carbon in fluvial bed sediments. Hydrobiologia 389: 153–167.
Sediment organic carbon
Source: ICRAM
Introduction
Carbon in marine sediments occurs as organic carbon (OC) intimately linked to the
metabolic processes and remains of plants and animals and as carbon contained within
biogenic and abiogenic carbonate minerals. Successful determination of sediment organic
carbon relies upon the separation of organic from inorganic carbon (IC). Several methods
of analysis have been employed for the separation of organic from inorganic forms of
25
carbon (reviewed in 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 presents difficulties. This is 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, reweighing of
sediment and analysis by CHN analyzer (Tung & Tanner, 2003).
Required
Falcon test-tube
Broad knife
Ruler (to measure sediment height)
Centrifuge (to extract interstitial water from the sediments)
Freeze-dryer (to lyophilize the sediments)
Mortar (to homogenize sediments)
Sieve with 200 µm mesh
Micro-analytical balance
Procedure
Sampling:
 Collect the sediment sample, by means of a corer device;
 Remove the upper 1 cm layer of the sediment, homogenize in a clean vessel, transfer
to a centrifuge tube and store on ice during transit to the laboratory
Sample processing:
 Samples are centrifuged to remove interstitial water and lyophilized (freeze-dried)
 Freeze-dried samples are gently pulverized using clean mortar and pestle and
subsequently sieved at 200 µm;
 6-10 mg of homogenized sediment are transferred to a special soft Silver cup (for
elemental analysis)
 Treat each sample by adding 20 µl of 1N HCl, to oxidize inorganic carbonate. If
effervescence continues, this step of the acidification procedure must be repeated;
 Wait for 10 minutes;
 Dry sample in 60°C oven for 10 minutes,
 Addition of 20 µl of HCl 1:1. If effervescence continues, this step of the acidification
procedure must be repeated;
 Wait for 10 minutes;
 Dry sample in 60°C oven for 10 minutes
 Addition of 20 µl of HCl 25%. If effervescence continues, this step of the
acidification procedure must be repeated;
 Wait for some minutes;
 Desiccate samples in oven, drying overnight at 60°C
26
Analysis:
Silver cups with samples are folded and carbon content is determined using a calibrated
carbon-hydrogen-nitrogen (CHN) Elemental Analyzer.
References
Tung, J. W. T. and Tanner, P. A., 2003. Instrumental determination of organic carbon in
marine sediments, Marine Chemistry, 80 (2-3): 161-170.
Verardo, D.J. 1990. Determination of organic carbon and nitrogen in marine sediments
using the Carlo Erba NA-1500 Analyzer. Deep-Sea Research 37 (1): 157-165.
Total nitrogen in sediment
Source: ICRAM
Introduction
The production of biodeposits (faeces and pseudofaeces) due to mussel cultivation, can
cause an increase in the total nitrogen concentrations in the sediment underneath mussel
lines (Chamberlain, 2002; Aleffi et al., in press). In a recent study (Christensen et al.,
2003) observed higher C/N ratios in surface sediments underneath mussel lines as
compared to a reference site. The same is true for sediments underlying intensive fish
farms, since the major fish species reared are carnivores that require high protein (= high
nitrogen content) diets.
Procedure
Sampling, sample processing and analysis are identical to those described for organic
carbon determination (above), but without the acidification steps.
References
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. Aquaculture 218: 567-588.
27
Inorganic and organic phosphorous in sediment
Source: Strickland and Parsons (1972) with modification by Aspila (1976)
Reagents
 Stock Ammonium Molybdate Solution
 Dissolve 30g of ammonium molybdate tetrahydrate in 1l of DI water
 Keep in a cool dark place



Stock Antimony Potassium Tartrate Solution
Dissolve 1.5g of antimony potassium tartrate in 1l of DI water
Keep in a cool dark place

10% Sulphuric acid



Ascorbic Acid Solution
Dissolve 60g of ascorbic acid in 1l of DI water
Keep in a cool dark place. Stable for about a month



Working Colour Reagent
Carefully mix 100mls of ammonium molybdate stock solution, 250mls of 10%
sulphuric acid 100mls of ascorbic acid and 50mls of antimony potassium tartrate
solution.
Keep in a cool dark place. Stable for about 6 hours

1M Hydrochloric acid


Potassium orthophosphate (KH2PO4) 10mM stock solution
Accurately weigh 136mg into 100mls of 1M hydrochloric acid.


Potassium orthophosphate standards
Prepare 0, 50, 100, 200, 400 and 800uM solutions by pipetting 0.5, 01.0, 2, 4 and
8mls of potassium orthophosphate standard into 1M hydrochloric acid and
making up to 100mls. This will give a calibration able to measure phosphorus
between 0 and 0.1% in a 0.3g dried sample aliquot
Procedure
Inorganic phosphorus
Weigh accurately between 0.2 and 0.3g of dried, homogenised and sieved (200 um)
sediment into a centrifuge tube and add 12mls of 1M HCl. Place on a shaking table for 16
hours and then centrifuge or filter. To 10mls of extract add 1ml or working colour reagent
and let stand for 20 min before measurement by spectrophotometer at 885nm. Prior to
measurement, calibrate the spectrophotometer with standards and the blank.
Calculate inorganic phosphorus content of sediment:
28
(u-molarity of extract) x (12/1000) x 31
1000000 x wt. of sample (g)
x
100%
Total phosphorus
Weigh accurately between 0.2 and 0.3g of dried, homogenised and sieved (200 um)
sediment into a crucible and ignite at 550 C for 1hour. After the crucible has cooled
empty contents into a centrifuge tube and add 12mls of 1M HCl. Repeat extraction and
measurement procedure of inorganic phosphorus determination.
Organic phosphorus content can then be calculated from:
Organic phosphorus = Total phosphorus – Inorganic phosphorus
References
Aspila, K.I. Agemain, H and Chau, A. 1976. A semi-automated method for the
determination of inorganic, organic and total phosphorus in sediments. Analyst, 101
187-197
Strickland J. and Parsons T. 1972. A practical handbook of seawater analysis. Bulletin
167 Fisheries Research Board of Canada, Ottawa
Water column (water quality) sampling
Selection of sampling stations
The same principles as described above, regarding sediment sampling, should dictate the
selection of water column sampling stations. Station positions should reflect gradients of
impact, normally aligned with the axis of predominant current and could consist of the
following distances from the farm: 0, 25, 50, 100, 200m and 1000m. Unlike the footprint
of fish farms that is often evident on the underlying sediments, the influence of
aquaculture activities on surrounding water quality is generally quite small and
measurable only within close proximity to the farm perimeter. It is noteworthy that in
highly dispersive areas, with fast currents, we may limit water column sampling to 2
stations – one in the vicinity in the farm and one at reference site, outside zone B,
because it is very difficult to detect a gradient in such highly dynamic environments.
Sampling dates should correspond to peak production and warmest water conditions in
order to capture maximal impact conditions. Samples should be taken along a diel or tidal
cycle (minimal sampling should include one at low water and another at high water in
tidal regimes) to include extreme conditions, e.g. DO sag at end of the night. Optimally,
should establish diel measurement of all variables at a minimum of 2 stations (e.g. one at
the farm site and second at anticipated intermediate effect station) within the area of
anticipated (from hydrographical data) effect and another at a distant reference station.
The water column will be sampled by CTD, and by bottle sampler at 3 depths; surface,
mid-depth and near bottom and Secchi disk depth will be determined at all stations. In
addition, the algal growth bioassay should be conducted at all possible study sites.
29
CTD vertical profiles at each station should include: salinity, temperature, sigma-t,
dissolved oxygen, chlorophyll, PAR (photosynthetically active radiation), light
transmission, turbidity (nephelometer).
Water samples will be collected using Niskin (or similar) bottles, and taken for dissolved
nutrients (including: ammonia, nitrate, nitrite, phosphate, silicate, total N, total P) and for
suspended particulate matter (including: total suspended solids (including POC, PON,
POP) and chlorophyll (for CTD fluorescence ground-truthing). For shellfish farm sites,
the phytoplankton community composition is extremely important and should be
characterized, where possible.
Secchi disk transparency
Source: Michigan Lake & Stream Associations, Inc
Background
The Secchi disk is used to measure how deep a person can see into the water. Even
though the Secchi disk measurement of water clarity is an approximate evaluation of the
transparency of water, it is used primarily for its simplicity. A more accurate
measurement of underwater irradiance can be made by the use of a photometer. Vertical
light penetration, as measured by Secchi disk, may also serve as a proxy for the
phytoplankton biomass in a water body.
The greatest value of the Secchi disk measurements occurs when readings at each station
are recorded and compared from week to week, month to month and season to season.
Several factors are involved in depth determination, including the eyesight of the viewer,
the time of day the readings are taken (midday- between 10:00 and 14:00 is preferred),
the reflectance of the disk, the color of the water, clay particles or other materials
suspended in the water, etc.
Equipment
Secchi disk - 8 inch diameter metal disk painted in alternate black and white quadrants
(see below) with waterproof tape or rope marked to 20cm resolution.
Procedure
The disk is lowered into the water by unwinding a waterproof tape or marked line (20cm
resolution) to which it is attached and until the observer loses sight of it. The disk is then
raised until it reappears. The depth of the water where the disk vanishes and reappears is
the Secchi disk reading. The depth level reading on the tape at the surface level of the
water is recorded.
In order that the Secchi Disk measurement be done to provide the greatest accuracy, the
following conditions should be met:
1. The same person should perform all readings since sharpness of vision varies
from person to person.
30
2. The reading should be taken on the same day of the week, if taking weekly
readings or at least not more than one day before or after the same day of the
week.
3. It is preferable that the measurement be taken between 10:00 a.m. and 2:00 p.m.
so that the light rays from the sky are at a similar angle each time the reading is
taken.
4. Avoid taking the measurement when the water is choppy or rough.
5. Readings should be taken on the shady side of the boat.
6. The reading should be taken at the same location each week.
Record:
a. Cloud cover (see Cloud Cover Field Guide below).
b. 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.
Secchi disk depth
procedure
31
Dissolved nutrients in seawater
Source: ICES 2004
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 compounds 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
following (including storage and pre-treatment) is Grasshoff (1976) “Methods of
Seawater Analysis”.
Sample handling
Special attention must be paid to possible nutrient sample contamination generated by the
boat or 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 nutrients 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 samples
The 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
32
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 new bottles, 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 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 abovementioned 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-treatment
Sea water contains microorganisms and other suspended matter of different composition.
In some cases, these particles bias the measurement of the nutrient 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
33
turbidity blank should be carried out by measuring light absorption of the sample before
adding the colour-forming reagents.
Appropriate chemical analytical methods
The 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
examination of large numbers of samples.
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 blank
Stock standard solutions should be prepared separately for each nutrient 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
34
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
photometric nutrient analyses. This bias can be avoided by measuring the sample before
addition of the colour reagent, or by filtration or centrifugation where this does not cause
contamination.
References
Grasshoff, K. 1976. Methods of seawater analysis. Verlag Chemie, Weinheim, New
York.
ICES. 2004. Chemical measurements in the Baltic Sea: Guidelines on quality assurance.
Ed. E. Lysiak-Pastuszak and M. Krysell. ICES Techniques in Marine Environmental
Sciences, No. 35. 149 pp.
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.
Nitrate/Nitrite in seawater (Cadmium reduction method)
(Lachat QuickChem method #31-107-04-1-C; http://www.lachatinstruments.com/)
Principle
Nitrate is quantitatively reduced to nitrite by passage of the sample through a copperized
cadmium column. The nitrite (reduced nitrate plus original nitrite) is then determined by
diazotizing with sulfanilamide followed by coupling with N-(1-naphthyl)
ethylenediamine dihydrochloride. The resulting solution has a magenta color, which is
detectable at 520 nm.
Procedure
Nitrite/nitrate will be determined on autoanalyser/flow injection analyser following the
standard colorimetric methods (Grasshoff et al. 1983). Samples containing high
concentrations of iron, copper or other metals may give low results. EDTA is added to the
buffer to reduce this interference.
OrthoPhosphate in seawater
(Lachat QuickChem method #31-115-01-1-G; http://www.lachatinstruments.com/)
Principle
Ammonium molybdate and antimony potassium tartrate react in an acid medium with
phosphate to form an antimony-phospho-molybdate complex. This complex is reduced
to an intensely blue-colored complex by ascorbic acid. The color produced is
proportional to the phosphate concentration in the sample. Though there is a density
difference between seawater and reagent water the bias is less than 2%.
35
Though the method is written for seawater and brackish water it is also applicable to nonsaline sample matrixes. The method is calibrated using standards prepared in deionized
water. Once calibrated, samples of varying salinities (0 to 35 ppt) may be analyzed. The
determination of background absorbance is necessary only for samples, which have color
absorbing at 880 nm.
Procedure
Orthophosphate will be determined manually or on autoanalyser/flow injection analyser
following standard colorimetric methods (Grasshoff et al. 1983).
Ammonium (Phenolate method) in seawater
(Lachat QuickChem Method #10-107-06-1-C; http://www.lachatinstruments.com/)
Principle
This method is based on the Berthelot reaction. Ammonia reacts with alkaline phenol,
then with sodium hypochlorite to form indophenol blue. Sodium nitroprusside is added to
enhance sensitivity. The absorbance of the reaction product is measured at 630 nm and is
directly proportional to the original ammonia concentration. Note: EDTA is added to the
sample in-line in order to prevent precipitation of calcium and magnesium ions. This
method is often adapted for use on an autoanalyzer, to allow for automated analysis of
multiple samples and replicates.
Total N & total P (persulfate digestion) in seawater
Source: Qualls (1989)
Introduction
This digest is applied to water samples to sweep the nitrogen from all N
compartments into nitrate and the phosphorus from all P compartments into
orthophosphate. The resulting digests are analyzed by manual or automated
colorimetry (e.g. Technicon autoanalyzer) for nitrate-N and orthophosphate-P.
This method is both safer and more effective than traditional Kjeldahl techniques.
It is based on the method of Koroleff (1983) as modified by Qualls (1989).
Equipment
1) Chemical autoclave or pressure cooker
2) 13x100 mm glass screw-cap culture tubes with teflon-lined caps. (Tubes are
acid-washed in 20% HCl and muffled at 500 degrees C for two hours. Caps are
acid-washed in 50% HCl)
3) autoclave-safe test tube racks
4) 100 ml, 200 ml or 500 ml acid-washed volumetric flask for oxidizing reagent
(depending on how much reagent is needed)
5) 500 ml acid-washed volumetric flask for 3.75M NaOH stock
6) 1000 ul and 5000 ul automatic pipetters
36
7) weigh boats and clean chemical spatula
Reagents
1) fresh deionized H2O
2) low-N potassium peroxydisulfate (e.g. Fisher P282-100)
3) boric acid (e.g. Baker 0084-01)
4) low-N NaOH if stock is needed
5) EPA-certified Nutrient 2 quality control digest standard
NaOH stock:
Place ~350 ml diH2O in 500 ml volumetric on stir plate. Add 75.0 g NaOH. Stir
to dissolve; remove stir bar and bring to volume. Cap with parafilm and invert to
mix. Allow to stand ~30 minutes and recheck volume.
Sample tube handling:
Wash all sample test tubes (70 mL etched glass tubes) with HOT phosphate free
soapy water and triplicate flush with dH2O. Acid wash the tubes and triplicate
rinse with DI H2O. After drying, store sample tubes capped until day of sample
collection.
Oxidizing reagent:
Place clean volumetric of appropriate size on stir plate; into it rinse in the
appropriate amounts of reagent from the table below with diH2O. Bring to about
80% of flask volume with diH2O and stir to dissolve; takes ~15 minutes on stir
plate (gentle warming may help.) When dissolved, remove stir bar and bring to
final volume with diH2O. Cap with parafilm and invert to mix. Allow to stand
~30 minutes and recheck volume.
100 ml
200 ml
250 ml
500 ml
persulfate
5.2 g
10.4 g
13 g
26 g
boric acid
3.12 g
6.24 g
7.8 g
15.6 g
NaOH stock
10 ml
20 ml
25 ml
50 ml
(This reagent may be stored 7 days at room temperature. Crystalizes when
refrigerated.)
EPA Nutrient-2:
10 ml concentrate from ampoule in 1000 ml diH2O (or 5 ml concentrate in 500
ml.) yields 5.00 mg/liter total nitrogen and 1.50 mg/liter total phosphorus.
(Digests for total nitrogen and total phosphorus are checked by digesting and
analyzing E.P.A. Nutrient-2 QC solutions formulated to challenge digestion
techniques alongside every batch of unknowns)
Procedure
1) bring samples to room temperature if chilled or frozen
37
2) make up fresh digest reagent, and NaOH stock if needed
3) obtain acid-washed, muffled digest tubes; label them.
4) for field samples (unknowns) and EPA2 (standard) samples:
 on first pass through the sample set, pipette 5 ml sample into each labeled
digest tube. Cap loosely to exclude dust.
 on second pass, pipette 1 ml oxidizing reagent into each digest tube. Cap
tightly and mix well (invert several times.)
5) For reagent blanks, pipette only 1 ml oxidizing reagent into tube and cap
tightly. (take care! Qualls (1989) states: "For low level samples the variability in
the reagent blanks determines the limit of detection, not the error associated with
the NO3 and PO4 analyses themselves.")
6) Place capped tubes in autoclave, 30 minutes on liquid cycle. (= 30 minutes on
"sterilize" in addition to all other cycle segments. If using pressure-cooker field
method, time 30 minutes after coming to canning temperature in addition to
warmup and cooldown times.)
7) After tubes are cool, add 5 ml diH2O to all reagent blank tubes so that the total
volume of liquid in these tubes is the same as in the others. (note, Qualls (1989):
"Since distilled or deionized water contains significant N, the dilution water [for
the blanks] is added after the digestion.")
8) Analyze digest-tube contents manually or with autoanalyzer using the nitratenitrite and orthophosphate manifolds.
Post-analysis calculations
1) Take the mean of the reagent blank determined values. Throw out any that are
>2 std. deviations above the mean (for nitrogen in particular this indicates that the
tube cap has cracked during the autoclave step and admitted atmospheric N to the
tube.)
2) To compensate for color absorption by the digest reagents, subtract the mean
reagent blank N and P values from the autoanalyzer determined values for each
unknown or EPA2 sample.
3) The effect of diluting the samples by the addition of digest reagents must be
reversed, using a dilution factor (df):
sample volume + reagent volume
df = -------------------------------------initial sample volume
In the case of the above procedure, where initial sample volume is 5 ml and
reagent volume is 1 ml,
5 ml sample + 1 ml reagent
df = ---------------------------------5 ml sample
=
1.2
Find the actual value of the undiluted sample by multiplying the determined value
(after reagent blank subtraction) by the df, as follows:
True analyte concentration
=
(raw determined value - rblank value)*(df)
38
Comments
1) Successful digests have pH in the range 5 to 8; incomplete digests are ~2. This
can be checked with wide-range pH paper. There is no reliable correlation
between final digest pH and the yellow color developed in some digests, so the
color cannot be used to spot incomplete digests.
2) Instead of using reagent blanks, it is possible to digest the calibration standards
(including water blanks, i.e. calibration standards of content zero), the D3
(recalibrant) and ref3 (reference check) and the W (baseline drift correction) cups.
Thus with digest reagent in both samples and calibants, the reagent's contribution
to total absorbance will be compensated for automatically. (This strategy is of
course useable only if all samples in the run are using the same reagent/diluent
ratio.)
It may be necessary to have different dilutions of the EPA2 QC standards for
nitrate and phosphate to get both into the optimum manifold range (e.g. if the PO4
manifold range is 0.2-1 ppm while the expected TP content of EPA2 is 1.5 ppm
PO4, the QC digests will be offscale for phosphate unless diluted.) Perform these
dilutions before digestion and then use the same reagent/diluent ratio for
everything.
References
D'Elia, C. F., P. A. Steudler, and N. Corwin. 1977. Determination of total nitrogen in
aqueous samples using persulfate digestion. Limnol. Oceanogr. 22, 760-764.
Koroleff, F. 1983. Simultaneous oxidation of nitrogen and phosphorus compounds by
persulfate. p.168-169. In K. Grasshoff, M. Eberhardt, and K. Kremling, eds., Methods
of Seawater Analysis. 2nd ed., Verlag Chemie, Weinheimer, FRG.
Koroleff, F.1983. Appendix A pp. 131-138. In The biogeochemical properties of
dissolved organic matter in a hardwood forest ecosystem: their influence on the
retention of nitrogen, phosphorus, and carbon. Ph.D. dissertation, University of
Georgia Institute of Ecology, Athens, Georgia, USA.
Langner, C. L. and P. F. Hendrix. 1982. Evaluation of a persulfate digestion method for
particulate nitrogen and phosphorus. Water Res. 16, 1451-1454.
Qualls, R. G. 1989. Determination of total nitrogen and phosphorus in water using
persulfate oxidation: a modification for small sample volumes using the method of
Particulate matter in seawater
Source: Plymouth Marine Laboratory and ICES (2004)
Introduction
This procedure addresses determination of Total Suspended Solids (TSS; which is also
known as Suspended Particulate Matter; SPM = total particulate matter, TPM),
Particulate Inorganic Matter (PIM) and Particulate Organic Matter (POM).
The particle size of 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
39
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 both major and trace elements.
Required
pre-washed, ashed and weighed GF/F 47mm filters, prepared as below, stored in clean
petri plates
clean forceps
100% acetone in wash bottle for washing off all surfaces that come in contact with filters
Freshly distilled water in wash bottle
Filtration manifold with filter holders for 47mm filters
0.5M Ammonium formate (31.5 g/l)
Dessicator
Drying oven
Muffle furnace
Concentrated HCl in Erlenmeyer or beaker for acid-fuming filters
Long (>20cm) forceps for holding filter over acid fumes
Fume hood for acid-fuming process
50mm (diameter) Petri plates
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.
Particulate Organic Matter and Total Particulate Matter determination :
a. Filter the required volume of homogenised material (see below)
b. Rinse filter twice with 10 ml of 0.5M Ammonium formate solution to remove salt and
then rinse with distilled water around margin of the filtration cup, having removed filter
head (do all of the above with pump running), and when dry, return filter to petrislide.
c. Oven dry filters (60°C for 2 days, 40°C for 1 week) and store in dessicator.
40
d. Weigh (from dessicator, to 5 places, as above, preferably with the same balance) for
total particulate matter (TPM).
e. Ash at 450°C in muffle furnace for > 4h
f. Weigh (from dessicator to 5 places, as above, preferably with the same balance) 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!
Reference
ICES. 2004. Chemical measurements in the Baltic Sea: Guidelines on quality assurance.
Ed. E. Lysiak-Pastuszak and M. Krysell. ICES Techniques in Marine Environmental
Sciences, No. 35. 149 pp.
Particulate organic carbon and nitrogen in seawater
Source: Plymouth Marine Laboratory and ICES (2004)
Required:
25mm filters prepared as below, stored in acid-washed Petri plates.
Access to CHN analyzer
Note: it is necessary to remove all organics and keep free from contamination using
gloves and covers, etc. The sample should be handled and transferred between containers
as little as possible to avoid contamination 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.
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.
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.
41
Storage of samples
Filters 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, preferably,
stored in a freezer and later dried in a drying oven at 60 °C for 30 min, cooled in a
dessicator and weighed prior to analysis.
Sample processing
The amount of sample (volume of water) required will depend on water quality.
Measures of chlorophyll 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 ml in times of
algal bloom or resuspension, but when seston levels are low, may need to filter up to 3
liters per sample (as has been done in eutrophic 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).
Examples, for mesotrophic waters (e.g. ~ 8 ug/l chl)
for TPM, PIM, POM
300ml on 47mm GF/F
(rinse with NH4 formate)
2 duplicate filters
for CHN
50ml on 25mm GF/F
(acid fumes)
2 duplicate filters
Therefore, for each sample, require: 2
2
2
for Chl a
10ml on 25mm GF/F
(no rinse)
2 duplicate filters
47mm GFFs (washed, ashed, weighed)
25mm GFFs (ashed only)
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 (see below)
must not be dried; instead, fold in half with material on inner side, and stored in freezer.
References
Grasshoff, K., Kremling, K., and Ehrhardt, M. (eds.) 1999. Methods of seawater analysis.
VCH, Weinheim, New York
ICES. 2004. Chemical measurements in the Baltic Sea: Guidelines on quality assurance.
Ed. E. Lysiak-Pastuszak and M. Krysell. ICES Techniques in Marine Environmental
Sciences, No. 35. 149 pp.
42
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.
Particulate organic phosphorus in seawater
Source: Marine Biogeochemistry - Practical Course in Biological Oceanography,
University of Kiel, Germany, 2005.
Organic phosphorus compounds are converted to orthophosphate (i.e. inorganic dissolved
phosphate) by cooking with potassium peroxydisulphate. The inorganic dissolved
phosphate is then measured colorimetrically, following Grasshoff (1983). During cooking
with peroxydisulphate, some chloride is oxidised to chlorine. Because it may interfere
with the measurement, this chlorine is then reduced by ascorbic acid (added in surplus,
before the addition of the mixed reagent; see below).
Interferences
The corresponding reaction with silicic acid (up to 200 μm l-1 Si) (see 4.5) does not occur
at a pH below 1.0. The reagents are designed for seawater in such a way, that the pH
adjusts itself around 1.0.
If the samples, already mixed with the reagents, are allowed to stand for more than 30
minutes AsO43- reacts slowly and a heteropoly acid forms. However, the arsenate content
in seawater amounts only to about 0.03 μm l-1 As.
Hydrogen sulphide in concentrations above 60 μmol l-1 H2S-S (= 2 mg S2- (sulphide))
disturbs the analysis. In such samples the sulphide has to be oxidized with bromine water
and the surplus bromine is driven out by airflow prior to analysis.
Calibration
From the stock solution (10 μmol ml-1 PO43--P) a calibration series of 0, 1 and 2 μmol l-1
PO43- is set up. With a pipette 0.1 or 0.2ml are transfered into a 1 litre measuring flask
and filled up to 1 litre with deionized water. For the 0 μmol l-1 PO43- standard only
deionized water is used.
The calibration series is treated as described in the chapter “Execution of the
Determination”.
Range: 0 – 10 μmol l-1 PO43--P (corresponding to 0 – 0,35 μmol/35 ml)
Precision: ± 0.02 μmol l-1 PO43--P (corresponding ± 0,0007 μmol/35 ml)
Procedure
A GF/F filter (blank or sample) is transferred to a 60ml Duran glass bottle and 35 ml of
deionized water are added to this. Next, 2ml potassium peroxidisulphate solution are
added, the bottle is closed and shaken. Samples are placed in an autoclave, or a household
pressure cooker filled 2 cm high with deionized water, and cooked for 30 min at 121 C
(at 15 psi), then cooled to room temperature. Next, 1 ml of ascorbic acid solution is added
to each sample. Shake and wait for 1 min before adding 1 ml of mixed reagent II. 12 ml
of this blue colored solution are transferred into centrifuge tubes and centrifuged at high
43
speed for 10 min. Following centrifugation, a sample of the cleared solution is transferred
to a cuvette and the absorption is measured at 882 nm against deionized water.
Reagents
4.5 M Sulphuric acid: Slowly add 250 ml concentrated H2SO4 (98%) to about 750 ml
deionized water and make up to a volume of 1000 ml.
(the mixture must be cooled – wear safety glasses!!!!)
Ammonium heptamolybdate: Dissolve 9.5 g (NH4)6Mo7O24*4H2O in about 50 ml
deionized water and fill up to 100 ml. Store in polyethylene bottles.
Potassium antimonyltartrate: Dissolve 3.25 g of potassium antimonyltartrate
K(SbO)C4H4O6*0.5H2O in 100 ml deionized water.
Mixed-reagent II: Slowly add 125 ml 4.5 M H2SO4 to 47 ml ammonium molybdate
solution, then 5.2 ml potassium antimonyltartrate solution and 73 ml deionized water
(total volume of 250ml)
Potassium peroxydisulphate: 30ml of 4.5 M sulphuric acid are given to 200 ml deionized
water. Then add 10 g of potassium peroxydisulphate (reagent remains stable during about
1 week)
Ascorbic acid: Dissolve 14 g ascorbic acid (C6H8O6) in 200 ml deionized water (at 4°C
durable for 30 days)
Stock solution: Dissolve 1.361 g dried potassium hydrogen phosphate (KH2PO4) in
deionized water and fill up to 1 000 ml (= 10 μmol ml-1)
Calculation of concentrations with a calibration series
Calibration series 0, 1 and 2 μmol l-1 (= 0,035 und 0,070 μmol/35 ml) 3–fold
determination
Examples
Ø EBl ~ 0.015 absorption of reagent blank
Ø ESt1 ~ 0.115 absorption of Standard 0,035 μmol / 35ml PO43—P
Ø ESt2 ~ 0.215 absorption of standard 0,070 μmol / 35 ml PO43—P
Chlorophyll a determination
Source: Plymouth Marine Laboratory
Required:
Clean water sampler, e.g. Niskin bottle
Filtration system with 25mm or 47mm filter holders
50mm Petri plates
47mm or 25mm GF/F filters
Filter forceps (flat)
Dispensor for 90% buffered acetone solution
Homogenizer for GF/F filters
90% buffered (MgCO3) acetone
Wash bottle with 90% buffered acetone
Glass bottle for acetone waste
Centrifuge tubes
44
Spatula
Lint-free tissues (e.g. Kimwipes) for cleaning cuvettes
HCl 1 N
Sampling
1. Filter water sample (e.g. 1000 ml) 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 labeled petri-dish and freeze until
required.
Chlorophyll extraction
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 refrigerate (4°C) for at
least 16, but not more than 24 hours 2. 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, 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. Scrape out any remaining bits of filter with spatula and add
these to centrifuge tube. Rinse homogeniser tube with remaining 6-8 ml 90% acetone and
add this to the centrifuge tube,. Cap centrifuge tube and store in refrigerator for between
16 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).
Note: Throughout storage and analysis of samples, exposure to light (especially
strong sunlight) should be avoided or at least minimised.
Fluorometric determination3
If possible, use a fluorometer (e.g. Turner Designs TD700) that does not require the acid
addition to. If not possible, measure sample fluorescence before and after addition of 2
drops (~100 ul) of 1N HCl to the cuvette.
Chlorophyll a (µg litre) = (FA (RB - RA)*Ve (ml))/Vf (ml)
FA = calibration factor of fluorometer (calculated as µg litre)
RB = fluorescence reading before addition of 1N HCL
RA = fluorescence reading after addition 1N HCL
2
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.
3 See Strickland J.D.Hand Parsons T.R. A Practical Handbook of Seawater Analysis. Fisheries Research
Board of Canada (Ottawa 1968) Bulletin 167.
45
Ve = Vol.of acetone extract (ml)
Vf = Vol. of sample filtered (ml)
Spectrophotometric determination
Use wavelength 750nm to measure turbidity and 663nm for chl a, using bandwidth of 1
nm. Ensure digital display knob is set on absorbance!
Measuring sequence: read absorbance at 750nm (E750), 663nm (E663), 750nm; add 2
drops HCl (leave 30 sec), read at 663nm (E663a)
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 HCL
E663a = abs. after addition of HCL
Quality Assurance
Sampling QA
• Keep the samples cool and in the dark.
• For chlorophyll a it is recommended that the sample is filtered immediately after
sampling or, at least, as soon as possible thereafter to avoid deposition of cells. If storage
is unavoidable, 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, and
evidence 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 is for equipment for measuring the whole spectrum (800–350 nm) for easier
checking of shifting of the chlorophyll peaks.
• Validate the spectrophotometer and the fluorometer at least once a year, or when
changes of the equipment are required.
• Calibrate the equipment with a certified reference material, if possible; use control
charts.
Reference
Strickland J.D.Hand Parsons T.R. 1968. A Practical Handbook of Seawater Analysis.
Fisheries Research Board of Canada (Ottawa 1968) Bulletin 167.
46
Bioassay studies to assess nutrient levels around aquaculture
Source: University of Crete (modification of Dalsgaard (2006) protocol)
Background
Release 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.
Required
Surface water
25 m mesh sieve/plankton net
Spectra/por 1 dialysis membrane
Collected from the control station of the site
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 wire
Nylon mesh bags to protect dialysis bags
Metal rod/plate
Rope
Weights (100g)
Buoys
Anchors
Bioassay setup
1. Cut the dialysis membrane into pieces of 30 cm length
2. 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 wire
4. Filter surface water from the control station of the site through a 25 m mesh sieve to
remove larger grazers
5. Dispense the filtered water into the dialysis bags (ca 600 ml/dialysis bag)
6. Close the dialysis bags with plastic coated metal wire
7. 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 bags
9. Using a rope and a metal plate hold together 5 replicate bags (Fig. 1)
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.
47
buoy
metal plate
dialysis bags
weight (optional)
mooring weight
Figure 1. Bioassay setup. Designed by the IMBC team participating in MedVeg project (based on
Dalsgaard 2006).
Analysis
1. Biomass of phytoplankton is measured as chlorophyll-a concentrations according to
standard protocols used for water column measurements
2. Before the analysis the volume of each dialysis bag must be recorded
References
Dalsgaard, T. and Krause-Jensen, D. 2006. Monitoring nutrient release from fish farms
with macroalgal and phytoplankton bioassays. Aquaculture 256:302-310.
Mura MP, Agusti S (1996) Growth rates of diatoms from coastal Antarctic waters
estimated by in situ dialysis incubation. Marine Ecology Progress Series 144: 237245
Mura MP, Agusti S, delGiorgio PA, Gasol JM, Vaque D, Duarte CM (1996) Losscontrolled phytoplankton production in nutrient-poor littoral waters of the NW
Mediterranean: In situ experimental evidence. Marine Ecology Progress Series 130:
213-219
48