Benthic fauna: collection and identification of macrobenthic invertebrates

Benthic fauna: collection and identification of macrobenthic invertebrates

Davide Tagliapietra and Marco Sigovini Istituto di Scienze Marine, Consiglio Nazionale delle Ricerche (CNR-ISMAR), Riva Sette Martiri 1364/a 30122, Venice, Italy

1. The benthos and its importance The community of organisms that live on, or in, the bottom of a water body is known as “benthos”. The term “benthos” (from ancient Greek âÝíèïò meaning “depth, depth of the sea, bottom”) was introduced by the eminent German naturalist and artist Ernst Haeckel (1834–1919), who also introduced the term “ecology”. The benthic community is complex. It includes a wide range of organisms from bacteria to plants (phytobenthos) and animals (zoobenthos) and from the different levels of the food web. Benthic animals are generally classified according to size: microbenthos 1.0 (or 0.5) mm and, sometimes, megabenthos > 10.0 mm. This exercise considers benthic animals, mostly invertebrates, lager than one millimetre, i.e. macrozoobenthos. Well-known groups of benthic animals are worms such as polychaetes and oligochaetes, molluscs such as bivalves and gastropods, and crustaceans such as amphipods and decapods. Benthic invertebrates can be differentiated by the position they occupy on or in bottom sediments: +

infauna - animals that live in sediments, almost all worms and bivalves belongs to this category, and

+

epifauna - organisms that live on the surface of bottom sediments; many crabs and gastropods are considered epifauna. Among the epifauna, animals can also be found living attached to hard surfaces, such as

the bricks and rocks of the banks or pilings. These are not so common in estuaries and lagoons. Epifauna includes also epiphytic invertebrates, i.e. organisms that live on the surface of submerged vegetation, such as many amphipods. Benthic invertebrates play an important role in transitional ecosystems, by filtering phytoplankton and then acting as a food source for larger organisms such as fish, thereby linking primary production with higher trophic levels. They also structure and oxygenate the bottom by NEAR Curriculum in Natural Environmental Science, 2010, Terre et Environnement, Vol. 88, 253–261, ISBN 2–940153–87–6

254

NEAR curriculum in natural environmental science

reworking sediments and play a fundamental role in breaking down organic material before bacterial remineralization. In addition, a number of benthic invertebrates, particularly clams, are consumed by humans and others, such as worms, are used for recreational purposes as fishing bait. Benthic communities are often used as biological indicators (e.g. the Water Framework Directive) because they can provide information on environmental conditions either due to the sensitivity of single species (indicator species) or because of some general feature that makes them integrate environmental signals over a long period of time. These features are: exposure to chemical contaminants often accumulated in the sediment; exposure to low dissolved oxygen levels (hypoxia/anoxia) that often occur near the bottom surface due to organic matter degradation; limited mobility that restricts their ability to avoid adverse conditions; taxonomic and functional diversity that make them suitable for the detection of different types and levels of stress. The composition, diversity and vigour of benthic communities also differ naturally according to the bioclimatic region, the type of ecosystem and the specific features of the habitat in which the organisms live, such as bottom water salinity, water renewal rate and sediment type. The purpose of this exercise is to introduce students to the technical procedures used in the collection and treatment of benthic samples in order to understand the actual difficulties of field work. The object of the study is the benthic macrofauna of coastal transitional ecosystems, i.e. estuaries, lagoons and coastal wetlands.

2. Sampling strategy The sampling strategy comprises a series of procedures aimed at selecting the most favourable sampling locations and determining the appropriate sampling density for accurate description and quantification of benthic assemblages in a given geographical area during a given period of time. The choice of strategy depends mainly on the distribution of the organisms belonging to the different species, on the characteristics and the number of habitats present in the investigated area and on the spatial and temporal variability of environmental parameters. It also depends on the feasibility, including the cost–effectiveness ratio. An in-depth discussion of the different sampling strategies is beyond the scope of this exercise. The main strategies for spatial allocation of sampling sites are: systematic sampling based on a regular distribution of the sampling sites over the whole investigated area, random sampling with sampling sites randomly distributed over the whole area, and stratified sampling. The latter is based on the identification of an area presenting reasonably homogeneous characteristics (stratum), for example vegetated bottom patches, bare substrate, etc. Sampling sites can be placed inside each stratum randomly or systematically or, in addition, the density of the sampling sites can vary inside every stratum. Once the sampling strategy and design has been defined according to the aim of the study, it is necessary to select the most suitable sampling instruments, determine the sample size and number of replicates, and select sampling techniques and treatments for samples.

Collection and identification of macrobenthic invertebrates

255

3. Field procedures Logistics are fundamental to all field activities. The choice of an adequate boat and equipment, the correct mooring, the ability to locate the planned sampling sites and the ability to return at any time, are all vital for good results. The other crucial requirement for effective teamwork is the achievement of harmonious working relationships. 3.1 Sample collection 3.1.1 Sampling devices Devices used to sample soft bottom macrobenthos basically comprise grabs, dredges, box-corers and hand nets. Grabs and corers are suitable for quantitative studies, i.e. when collection of a defined amount of sediment is required. These devices allow good reproducibility and reliable replicates of the samples. A variety of grabs and box corer is available from specialized suppliers. The choice of the most suitable device depends on various factors including operational conditions, substrate and physical characteristics of the investigated habitat. Researchers often modify sampling devices to meet local requirements. For use in wetlands there is a general convergence towards devices of moderate weight, in order to allow adequate penetration into the sediment without being too heavy to be hand-operated. The most widely used grabs are the Van Veen and the Ekman-Birge (the latter is often referred to as a box-corer because of its shape) (Figure 1). The Van Veen grab is easily operated by a rope and, therefore, it is suitable for relatively deep waters but has the disadvantage that it takes an uneven mouth-shaped bite of the sediment. Conversely, the Ekman-Birge grab is operated usually by a handle and gives a perfect box-shaped sample, which is very useful for volumetric analysis. This device preserves the substrate stratification and is therefore suitable for research on the vertical distribution of organisms. The area sampled by a device is, in general, a compromise between ease of handling and the need to get representative samples; a high number of relatively small samples often yield better information than a very small number of large samples. A surface area of about 200–600 cm2 is typical for grabs deployed in transitional environments. Replicate samples are usually needed because of the variability in spatial distribution of macrobenthic organisms. As a general rule 3–5 replicates are considered adequate to represent the benthic community of an estuarine site, but the correct number of replicates can be obtained only after analysis of species–area relationships in a given assemblage. A sampling device must recover relatively undisturbed samples to a depth sufficient to collect the majority of the burrowing organisms dwelling in the deeper layers of the soft bottom. In coastal wetlands a sample depth of about 15–20 cm is usually adequate. The Ekman-Birge grab, 15×15×15 cm in size, is particularly suitable when used with small boats for sampling on muddy bottoms in shallow waters (Figure 2). Other semi-quantitative samplers include Surber samplers that have a square-shaped frame with a long handle and an attached net, usually with 500 µm mesh size. They are usually

256


Figure 1 Popular samplers for soft-bottom sediments: the Van Veen (left) and the Ekman-Birge (right) grabs

used in river sampling, but are also used in coastal transitional environments for epiphytic organisms only (i.e. organisms living on submerged vegetation). In some cases “suction samplers” may be used for collecting soft bottom macrobenthos. These devices are tubes that suck organisms and bottom sediments into a net, similar to the operation of a vacuum cleaner. 3.1.2 Sieving Samples are sieved in order to remove fine sediments and any other extraneous material. Once on board the boat, the grab is opened above a plastic bucket and the sample gently removed. Before sieving the sample should be described and notes on surface characteristics, individual density, occurrence of organic detritus, etc. recorded in a sampling form. When clay sediments are present, which is common in estuaries, it is advisable to break up the sediment in water inside the bucket by adding filtered seawater and stirring gently. Filtered water is used to avoid the introduction of unrelated small organisms. The sample is then sieved; water is sprinkled directly onto the sample with a low-pressure nozzle in order to prevent any damage to animals. If the boat is too small or time is short, the samples can be kept in watertight plastic bags in a thermally isolated container and, during the warm season, cooled with icepacks. The delicate process of sieving should be performed very carefully in order to avoid any damage to the fragile organisms and to ensure that all animal present in the sample are collected. In order to separate macrofauna, a sieve of 1 mm or 0.5 mm mesh is used. A 1 mm mesh is preferable when the sediment contains a large amount of detritus, as often happens in wetlands and estuarine environments, in order to prevent clogging of the sieve. In any case the sieve must have an adequate surface to avoid clogging. Aquatic vegetation present in the sample should be cautiously removed from the surface of the sample, rinsed apart and the resulting water sieved. Small circular sieves are suitable for manual sieving, whereas bigger sieves are suitable for fitting into sieving desks. All material retained on the sieve, including organisms, shell fragments, vegetal debris and coarse sediment grains, are transferred to appropriate containers. The material is removed from the


257

Figure 2 Biologist using an Ekman-Birge grab in the field (Porto Marghera Lagoon of Venice).

sieve using a water jet and conveyed through a funnel into a fine mesh bag (a nylon sock) fixed to the outlet of the funnel. The bag is then put into a suitable plastic container and labelled. Containers must be labelled both internally and externally; the external label can be written with a permanent marker, the internal label can be made of tracing paper, written in lead pencil or Indian ink. The labels must record: the station code, the sample code, sampling date and split number for any sampling replicates. The split number is the partial number of a series; e.g. 2/4 refers to the second replicate of a series of four. Labelling should be performed by two operators, and recorded in special files and checked by the field supervisor. 3.1.3 Fixation and preservation Before fixation, the sample contained in the bag should be placed in a narcotic and relaxant solution (e.g. a solution of 7% magnesium chloride or 10–15% ethanol) to relax the organisms and avoid unnecessary suffering. Fixation of a biological sample stops the post-mortem degeneration of tissue induced by autolysis and prevents microbial attack, thus preserving the structure as unchanged as possible. Fixatives denature proteins, resulting in the hardening of tissues.

258


The sieved material is fixed as a whole in the plastic container. The volume of the fixative should be approximately three times the volume of the sample. The presence of considerable quantities of organic matter requires a larger quantity of fixative. The fixation of organisms may be achieved within a couple of days. The most common fixative for benthic organisms is a 10% formalin solution (or 4% formaldehyde). Formalin is a commercial aqueous solution of 40% formaldehyde. Formalin is an acid, therefore it should be buffered in order to avoid the dissolution of calcareous parts of the organisms. Extreme caution should be used in the manipulation of formalin because formaldehyde is toxic and carcinogenic. Less toxic fixatives are available as an alternative; for example an alcohol such as denaturized ethanol can be used. It does not perform as well as a fixative but it is much less toxic than formalin. After a few days, the samples are fixed and can be removed from the fixing solution, rinsed and placed in a preserving solution. The most common preservative is an aqueous solution of ethanol composed of 70% ethanol and 5% glycerin. Some researchers find it useful to stain the sample to accelerate the sorting procedure. One of the most common stains is Rose Bengal, which should be used carefully and sparingly because it is considered carcinogenic. In some cases Rose Bengal stains part of the detritus and it is more of a nuisance than a help. Samples should be accompanied by a “Chain-of-custody” form, which is a special form indicating the origin, delivery date, type of transport and the names of the persons in charge of loading, transporting and unloading the samples. The record must enable materials and persons in charge to be traced.

4. Laboratory procedures 4.1 Sorting Sorting consists of picking up from the sieved material all the animals that were alive at the moment of the sampling. Sorting procedures are performed under fume hoods to prevent inhalation of vapours of residual toxic substances. Large samples can be subdivided into sub-samples of roughly equal size that can be sorted more comfortably. The sub-samples should be placed in different jars with preserving solution and labelled. A small quantity of unsorted material is placed on a tray for an initial general sorting for larger organisms with the help of a magnifying lens. Shell fragments, vegetal debris or coarse detritus in the sample should be rinsed in a separate container and checked for the presence of invertebrates. Large organisms are placed immediately in appropriate containers making sure that no other smaller animals are attached to their bodies. Fine sorting is performed under a dissection microscope. During this phase a small quantity of the sample is spread onto a Petri dish and carefully examined to identify the organisms. Organisms are picked up and placed in different containers according to the main taxonomic groups, usually polychaetes, other worms (oligochaetes, nematodes, nemertines, etc.), bivalves, gastropods, amphipods, other crustaceans, insects, cnidarians, sponges and other


259

animals (a.c. = animalia cetera). Containers are labelled inside and outside. Cross-checking between operators is important during sorting because it can be very tiring and there is a high risk of error resulting from loss of concentration. 4.2 Identification procedures Once sorted, animals are identified to the taxonomic level required by the investigation. For monitoring purposes, the level of taxonomic identification required is sometimes the family or even higher level, but often the species level is required. The instruments used in identification are a dissecting stereomicroscope coupled with a compound microscope when observation of fine details is needed. Identification is done with a help of identification keys. There are keys for any major group of organisms, for example, gastropods, bivalves, amphipods etc. The keys are structured with a series of two choices to be made about the anatomy of an animal (“dichotomous,” two branches), and the answers progressively reduce possible identification choices until a single name is left. The use of identification keys generally requires more than a basic understanding and thorough training is required for the identification of anatomical characteristics. For correct identification, accurate analytical keys for the geographic region from which the samples were taken should be used. To catalogue species correctly it is strongly recommended that the international checklists of species, e.g. the European Register of Marine Species (ERMS) or Integrated Taxonomic Information System (ITIS), or national checklists are consulted. Usually each laboratory organizes its own “reference collection” consisting of a number of specimens identified by experts that serve as a reference to check the correct identification of an organism. A high level of expertise is often required to achieve complete identification of the benthic fauna of a coastal wetland. Experts also need to confirm identification from their laboratory with those of colleagues and specialists of different taxonomic groups and may also need to attend training courses. The importance of well-trained specialists in taxonomic identification cannot be overemphasized.

4.3 Determination of key descriptors The main descriptors of a benthic community that can be obtained from the samples are: a) Species richness (the number of species), b) the Abundance (the counts of individuals for every species) and c) the Biomass. The last two parameters are also called “importancies” because they give the “influence” of the different species in the numerical analysis. Other descriptors that can be obtained during this phase are the size-spectra of the different assemblages. Organisms in the same sample that belong to the same species must be counted and stored together in a test tube containing preservative solution before determination of their biomass. Counts of different species found in a sample are reported in a datasheet and later transferred to an electronic spreadsheet.

260


Beside the numerical abundance of the organisms, another important parameter is their biomass i.e. the weight of the living matter. The water content of organisms is highly variable and it is therefore preferable to estimate biomass after drying (dry weight) at 60 °C for 24 hours. The biomass of organisms is the sum of soft and mineralized parts (the latter is much less metabolically active than the former). A convenient method for obtaining an estimate of the metabolically-active biomass is the calculation of the so-called Ash-Free-Dry-Weight (AFDW), which is the value of the dry weight minus the weight of ash. Ash is obtained by heating the sample in a muffle furnace at 450 °C for about 8 hours.

5. Sources of information BLM/USU National Aquatic Monitoring Center Standard Sample Sorting Procedures, National Aquatic Monitoring Center, a.k.a. The BugLab, Department of Fisheries and Wildlife, Utah State University, Logan, Utah USA 84322-52 http://www.usu.edu/buglab/sortproc.pdf [Accessed 10 February 2009]. Barnes, R.S.K. 1994 The Brackish-Water Fauna of Northwestern Europe: A Guide to Brackish-Water Habitats, Ecology and Macrofauna for Field Workers, Naturalists, and Students. Cambridge University Press, Cambridge, UK. Castelli, A., Lardicci, C. and Tagliapietra, D. 2004 Soft-bottom macrobenthos, Chapter 4, In: M.C. Gambi and M. Dappiano [Eds.] Mediterranean Marine Benthos: A Manual of Methods For Its Sampling and Study, Biologia Marina Mediterranea (SIBM-APAT-ICRAM), 11(Suppl. 1), 99-131. Cuffney, T.F., Gurtz, M.E. and Meador, M.R. 1993 Methods for collecting benthic invertebrate samples as part of the National Water-Quality Assessment Program. United States Geological Survey Open-File Report, 93-406. Davies, J., Baxter, J., Bradley, M., Connor, D., Khan, J., Murray, E., Sanderson, W., Turnbull, C., and Vincent, M. [Eds] 2001 Marine Monitoring Handbook, Joint Nature Conservation Committee. Dybern, B.I., Ackefors, H. and Elmgren R. [Eds)] 1976 Recommendations on methods for marine biological studies in the Baltic Sea. The Baltic Marine Biologists Publication, 1. Holme, N.A. and Mc Intyre, A.D. [Eds] 1984 Methods for the study of marine benthos. Blackwell Scientific Publications, Oxford UK. International Council for the Exploration of the Sea (ICES) 2004 Report of the ICES/OSPAR Steering Group on Quality Assurance of Biological Measurements in the Baltic Sea. ISO 16665: 2005 (E). Water quality – Guidelines for quantitative sampling and sample processing of marine soft-bottom macrofauna. International Organization for Standardization, Geneva. Manual for Marine Monitoring in the COMBINE Programme of HELCOM, 2003. Part C. Programme for monitoring of eutrophication and its effects. Annex C-8 Soft bottom macrozoobenthos. http://www.helcom.fi/groups/monas/CombineManual/PartC/en_GB/main/ [Accessed 10 February 2009] Moulton, S.R.Ii, Carter, J.L., Grotheer, S.A., Cufney, T.F. and Short, T.M. 2000 Methods of analysis by the U.S. Geological Survey National Water Quality Laboratory processing, taxonomy, and quality control of benthic macroinvertebrate samples. United States Geological Survey Open-File Report 00-212. Rumohr, H. 1999 Soft bottom macrofauna: Collection, treatment, and quality assurance of samples (Revision of No. 8). ICES Techniques in Marine Env. Sciences, 27, Copenhagen DK Russel, D. 2001 USEPA Region 3 Quality Assurance Guidelines for Sorting and Identifying Marine Benthos. DCN: R3-QA500, DCN: R3-QA500-: R3-QA501 USA


261

SCBFMC (Southern California Bight Field Methods Committee) 2002 Field operations manual for marine water-column, benthic, and trawl monitoring in Southern California. Southern California Coastal Water Research Project Authority (SCCWRP) USA Strobel, C.J., Kemm, D.J., Lobring, L.B., Eichelberger, J.W., Alford-Stevens, A., Potter, B.B., Thomas, R.F., Lazorchak, J.M., Collins, G.B. and Graves R.L. 1995 Environmental Monitoring and Assessment Program (EMAP)- Estuaries: Laboratory Methods Manual, Vol.1 Biological and Physical Analyses. Office of Research and Development, U.S. Environmental Protection Agency, Narragansett, RI USA. Todorova, V. and Konsulova, Ts. 2005 Manual for quantitative sampling and sample treatment of marine soft-bottom macrozoobenthos, IO-BAS-Varna, Bulgaria