Macrobenthos procedures
Macrobenthos Procedures
Assorted Sources
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.
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 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 soft-bodied taxa that are difficult to identify (e.g., some polychaetes) and smallest for taxa encased in a hard enclosure such as a calcareous shell (e.g., most molluscs) or an exoskeleton (e.g., crustaceans), particularly if the hard parts are the primary taxonomic characters used for identification.
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
*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 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).
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).
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 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).
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.
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.
