Study methods

Collecting and sample processing

As with most invertebrates, searching for polychaetes in the field is not productive except for a few large species that are readily targetted. If representative samples of a polychaete fauna are required it is necessary to collect a sample of the substrate (eg sand, mud, algae) and sort the worms and other organisms from the sample in the laboratory. This should be done using a dissecting microscope to ensure all specimens in the sample are found. Collecting should be carried out in an environmentally responsible fashion, and permits are now required for most such activities.

On rocky shores, polychaetes occur in crevices, under boulders, and associated with algal holdfasts, tufted coralline algae, sponges and ascidians. Serpulids which live in calcareous tubes are firmly attached to hard substrates such as rocks, pilings and mollusc shells. An oyster knife is useful for prising off algal holdfasts and tufted algae which should then be placed immediately in a container with seawater and gently agitated and many species will swim out of the habitat. Remember to turn back any boulders which you have turned over.

Polychaetes which live on sandy and muddy shores can be collected using a spade or a corer (a cylinder that is pushed into the sediment, retrieving a "core"); the sediment is gently sieved and the individual species picked off the sieve with forceps. Many tubiculous species betray their presence in the sediment by the tops of their tubes or by mounds of faecal pellets such as the arenicolids. Free living species can rapidly burrow and some tubiculous species retreat rapidly deep into their burrows, so dig quickly.

Collecting from shallow subtidal habitats can be carried out by using SCUBA, and either using hand operated corers or collecting particular habitats such as algal holdfasts, seagrass beds, algal clumps, sponges, ascidiians, clumps of mussels etc., which should then be placed immediately into bags and sealed, before inquisitive fish arrive. Polythene bags are difficult to use underwater and a 0.5 mm (or smaller) nylon mesh bag with a drawstring closure may be easier to fill with the sample, and they are also stronger. Small airlifts operated from a SCUBA bottle can also be used to vacuum soft sediments and seagrass beds. Man made structures such as wharf piles and jetties are often colonised by a diverse encrusting community which may contain large numbers of serpulid and sabellid polychaetes as well as other species. Quantitative studies of these communities often involves using quadrats and scraping all the fauna within that quadrat into a polythene bag. Photographing the quadrat may also be useful in estimating the % cover by the various organisms, although individuals will need to be examined in order to determine species.

In coral reef waters, dead coral substrate contains large numbers of polychaetes representing many families, and the colonies should be chipped off with a hammer and chisel and placed immediately in a bag and sealed.

Myzostomes can be collected from the arms of crinoids, which should also be collected to determine the host species on which they are living. Other specialised habitats for polychaetes include the undersurface of holothurians.

Sample notes on locality, depth and habitat should be made on an underwater slate and later transferred to the labels for each sample.

In many cases, SCUBA diving is not an option and benthic samples need to be collected by the use of a grab or corer operated from a boat. The use of a dredge is not highly recommended, although some polychaetes may be collected but more often the tops of tubes only. Grabs or corers when they are returned to the surface need to be processed immediately. Grab samples should be emptied onto a sieve and washed gently to minimise damage to the worms, such as loss of palps and scales and the animals removed with forceps. Core samples and grab samples which cannot be sieved should be placed in fine mesh bags and placed in a large container of 7% neutralised formalin and gently agitated to ensure that the preservative penetrates the entire sample. To facilitate sorting, Rose Bengal can be added to the sample which stains all living material pink. Other organisms such as sponges and mollusc shells may provide habitats for polychaetes and these should be carefully checked.

For investigation of the spatial distribution distribution of polychaetes within a core, such cores should be rapidly frozen and then cut into slices and preserved.

All samples should be labelled using water proof paper and ink, when the entire sample is fixed for later sorting, such labels should be placed within small plastic bags and sealed within the larger bag. Details of the exact position of each station should be recorded as well as depth.

Some polychaete families are entirely pelagic and can be collected using plankton nets. Many families have pelagic larvae and others have a modified swimming stage - an epitoke or heteronereid associated with reproduction and these can be collected at restricted times using plankton nets and they are strongly attracted to strong lights. All these pelagic families and reproductive stages are very fragile and the worms easily fragment.

Relaxation, Fixation & Preservation
A variety of narcotics have been successfully used to immobilise polychaetes, either in preparation for fixation, or prior to photography. These include:

• Magnesium chloride is an effective narcotic for marine invertebrates at a concentration of about 7% in freshwater.
• Menthol crystals scattered on the surface of a seawater dish will narcotise invertebrates, but it may take 12 hours.
• MS222 can be used successfully to narcotise polychaetes. Advantages include quick recovery from the anaesthetic and effective at very low concentrations. Also there is no powder/refractive index change, an important consideration if photography is involved. A disadvantage is that MS222 is toxic and requires appropriate safety procedures in handling it.
• Phenoxyethanol is a useful narcotic, although it too is toxic. Mix with freshwater then dilute with seawater containing your animals (final concentration should be less than 1%). Usually works within half an hour. It seems to lose its potency over time if mixed directly with seawater.

George, J.D. and Hartmann-Schröder, G. 1985. Polychaetes: British Amphinomida, Spintherida & Eunicida. Keys and Notes for the Identification of the Species. Synopses Of The British Fauna New Series No. 32. E. J. Brill & Dr W. Backhuys: London 221 pp.
General overview of the methods used in studying polychaetes.

Gibbs, P.E. 1976.Fixation and preservation of planktonic polychaeta. pp. 279-280 in Steedman, H.F. (ed.), Monographs On Oceanographic Methodology, Vol. 4. Zooplankton Fixation and Preservation. . Illus. Unesco Press: Paris, France. (Available from Unipub: New York, N.Y., USA.).
Good for methods specific to the study of pelagic polychaetes.

Mackie, A.S.Y. 1994. Collecting and preserving polychaetes. Polychaete Research 16, 7-9.
Good general overview of the methods used in studying polychaetes.

Westheide, W. 1990. Polychaetes: Interstitial Families.. Universal Book Services: Oegstgeest. 152 pp.
Overview of methods specific to the study of meiofaunal polychaetes.