Collecting marine invertebrates

Page Contents

Leslie Harris, 2016

Why live-sorting

This guide is primarily aimed at live-collecting in the field and maintaining the collection alive until arrival at the laboratory for live-sorting and preservation.

Advantages of live sorting for the small and squishy:

The disadvantage of live-sorting in the lab is that some organisms (some small crustaceans, for example) may not survive long enough. The best way to preserve DNA is to place a tissue sample directly into 95% ethanol (or RNAlater®) immediately in the field — DNA recovery starts to decline once animals are dying. If the goal is preservation of DNA, consider preserving some or all of the collection in 95% ethanol immediately in the field.


Do not add fresh water

Under no circumstances should an unfixed bulk sample be washed in fresh water before preservation if anything but crustaceans, molluscs, and echinoderms are wanted. The abrupt change in osmotic pressure can and will cause severe cell damage in soft-bodied animals making them unfit for both morphological study and DNA work.

Although adding a small amount of ethanol or fresh water to an unsorted sample will provoke many animals into moving away from the substrate it can also damage them. Other animals — soft-bodied ones in particular — react to these irritants by going deeper into their substrate which makes them harder to find. Again, for multi-phylum samples this is not recommended.

Keep certain taxa separate

Certain taxa should be kept separate from other samples, and should be processed first upon arrival in the lab.

Sponges in particular can rapidly deteriorate and release noxious compounds; their cut or torn surfaces will also release compounds.

Encrusting colonial tunicates like Botryllus and Didemnium can quickly go bad.

Solitary and large colonial tunicates are usually harmless but they remove large quantities of oxygen from the water which is a problem with inadequate aeration.

The brown alga genus Desmarestia (which may be collected for its associated invertebrates) contains sulfides lethal to other animals.

Transporting and maintaining live samples

Use battery-operated air pumps and frozen cold packs (or soft-drink bottles filled with water and frozen) to keep the water aerated and cold and the unsorted samples alive.

Don’t overfill the temporary container with sample: too much biomass to too little water will result in poor samples even with aeration and cooling.

Under optimal conditions, unsorted live samples can stay in good condition for several days.


Sort the animals by phylum, class, order, morphospecies — whatever is desired for initial processing. Use the lowest impact method to capture the animal. Spoons for larger animals, pipettes and paintbrushes for smaller ones are much better than forceps. It is always best to put large, medium, and small size groups into separate dishes. Jumbling the sizes together in one dish results in smaller individuals being lost or eaten. Take extra care to keep sediment and floc out of the animal dishes. These accumulate on appendages, obscuring features and acting as substrates for salt crystal formation during preservation. Some animals actively take up sediments as camouflage while worms and some peracarids start making tubes with it, requiring much extra effort to clean them for photography, effort which could have been avoided if the water stayed clean in the first place.

Process sponges and tunicates first, since they can rapidly deteriorate and release noxious compounds.

Sponge examination: Wear gloves. Shake sponges in a container to detach vagile animals from the surface or visually inspect them. Next, gently tear the sponge to see if there are animals living within or under the structure. Polychaetes, flatworms, ophiuroids, small clams, and peracarids are often found inside sponge. If any are seen, the sponge should be examined with a magnifier or dissecting scope to facilitate picking out the animals without damaging them. This is especially important in the case of worms and ophiuroids, which end up in pieces if the sponge is carelessly torn.

Tunicate examination: Shake tunicates in a container to detach vagile animals from the surface or visually inspect them. Tunicate bases of both solitary and colonial forms should be examined under a magnifier or dissecting scope for epifauna. They may harbor amphipods and shrimp inside their bodies; depending on whether the commensals are inside the tunicate or inside a siphon you can get them out by squeezing or tearing.

Algae-associated invertebrates: Most temperate algae will survive in aerated cold water without problems or affecting other organisms, but, like tunicates, will use up dissolved oxygen. The one exception is the brown alga genus Desmarestia which contains sulfides lethal to other animals. Algae samples should be gently washed several times in a large amount of seawater to remove free-living individuals, then examine the fronds to be certain clinging cryptic organisms aren’t missed, and finally cut the fronds off a little above holdfasts and discard the fronds. Swirl the container containing the wash to get the light organisms up into the water and gently pour the water into a sieve. Rinse the animals off the sieve and into a large dish with seawater. Examine the holdfasts with a magnifier or dissecting scope for microfauna.

Holdfasts, Mytilus and Pollicipes colonies, clumps of Phragmatopoma, oysters, etc. all form complex three-dimensional living spaces perfect for small organisms. Gently separate the colonies in a container under water and shake to remove animals; the Phragmatopoma can be gently crushed. Rapidly swirl the container to get the lighter animals up into the water and gently pour the water into a sieve. Rinse the animals off the sieve and into a large dish with seawater. Repeat at least twice. The residue at the bottom of the container should be spread out in a tray with ample water to check for animals too heavy to float. This grunge — or a fraction of it — should also be examined with a dissecting scope as polychaetes, acoels, flatworms, nemerteans, sipunculids, tiny snails and clams, peracarids, and echinoderms small enough to fit into micro-crevices will still be in it. Many of these tiny animals are adults not found otherwise, and can add a considerable number of species to the final list.


In general, soft-bodied animals require extra care in order to produce good specimens suitable for morphological identification. Even some animals with exoskeletons like ophiuroids need to be relaxed as well to prevent the arms from breaking. It’s useful to relax bivalves and gastropods so the valves/opercula are open and fixative can penetrate the body.

Preservation of DNA is best done with live tissue (by subsampling an individual’s tissue or preserving an entire individual), and should precede relaxation — some relaxants may damage DNA. Sampling for DNA must be done before any formalin-based fixatives are applied.

Sponges can be dropped directly into fixative. If the whole sponge is not preserved a subsample consisting of both outer and inner regions is acceptable to specialists.

Tunicates must be completely relaxed with the atrial apertures open before fixation, as internal anatomy is crucial for accurate identification. Subsamples for DNA should be taken before relaxation, or taken from a second animal. Relaxation can be started in the field by placing them into seawater-filled whirl-paks to which a few crystals of menthol or several drops of saturated menthol in 95% ethanol solution has been added. Magnesium chloride can be used instead. Narcotization can take up to several hours before the animals no longer react to a probe. Quickly rinse them in seawater to remove extra crystals and put into 10% buffered seawater formalin.

Cnidarians: Anemones need to be relaxed before fixation. Subsamples for DNA should be taken before relaxation, or taken from a second animal. Formalin is preferred for fixation, as internal features are important for morphological identification.

Crustaceans can go directly into 95% ethanol. If they will be photographed, they need to be relaxed with 7.5% magnesium chloride or killed by putting them into a freezer. This keeps the color intact and the limbs supple so they can be cleaned and posed.

Polychaetes need to be relaxed, especially if they’re going into 95% ethanol. Subsamples for DNA should be taken before relaxation, or taken from a second animal. It’s best to have several individuals per species, with some being put into ethanol for DNA and others going into 5–10% buffered seawater formalin. If there’s only one specimen, it should first be relaxed with 7.5% magnesium chloride, then a small median section can be removed for DNA while the rest goes into formalin. Animals of different families, species, and sizes within species react differently to 7.5% magnesium chloride. Large nereids and thick-scaled polynoids such as Halosynda can be put directly into the relaxant without ill effects. Phyllodocids and syllids react violently to magnesium chloride unless it is slowly and carefully dripped into the dish. Ask Leslie Harris ( for advice if you end up dealing with polychaetes.

Nemerteans and flatworms: Treat these gently to avoid breakage. Use a spoon or a paintbrush to pick up flatworms. They must be thoroughly relaxed and have their color documented before preservation. In the absence of unique color patterns (which may actually belong to sibling species) identification requires examination of internal characters which are difficult and sometimes impossible to see if the worms contract and contort during preservation. Some specialists will not accept specimens that were not relaxed prior to fixation (Jon Noremburg at the Smithsonian is one). Keep checking them during the relaxation process as both types can seem fine one minute and literally start dissolving the next.

Nemerteans can be dropped directly into 7.5% magnesium chloride or add the solution slowly to seawater until a 1:1 mix is attained. Subsamples for DNA should be taken before relaxation, or taken from a second animal. Both head and tail features are necessary for ID so take a median section for DNA once the animal is relaxed. Formalin is best for fixation, as it preserves internal characters better, followed by a rinse in fresh water before being placed in 70% ethanol.

Flatworm preservation ideally requires not only meticulous care but also frozen formalin. Subsamples for DNA should be taken before relaxation, or taken from a second animal. The full fixation procedure is described below. An easier method is to put the flatworm into a 15 mm petri dish lid with a few drops of water, then put the inverted slightly smaller bottom on top of it. This keeps the worm flat while the lower dish is quickly flooded with 5-10% formalin or 95% ethanol. Others use 2 slides banded together and again, quick flooding with fixative. Use a second worm for DNA or snip a posterior piece off before preservation. They are difficult to relax, often disintergrating before total narcotization is achieved.

Polyclad fixative

The following recommendation on fixation of polyclad flatworms is quoted from: Newman, L, Cannon, L. 2003. Marine flatworms: the world of polyclads. CSIRO Publishing, Australia.

To make 1 litre of polyclad fixative you need 100 ml of concentrated formaldehyde which you dissolve in 900 ml of seawater, and then there are some additional ingredients which may prove to be optional, especially for acotyleans or temperate species. These ingredients are 2 g of anhydrous calcium acetate, 45 ml of propylene glycol (propane 1,2-diol) and 5 ml of propylene phenoxetol (phenoxy-propan-2-ol). These last three additives are especially valuable for preserving the delicate outer layers and for this we are indebted to the work published in 1991 on terrestrial flatworms by Leigh Winsor.

The procedure

Pour the fixative solution into a fixing container (plastic take-away containers are excellent) to cover the bottom to about 1-2 cm and freeze solid. Note that this method only works if the fixative solution is frozen! The procedure works because the normal reaction of worms to stress and danger is to flatten out into a thin sheet against the substrate they are crawling on. So the idea is to get the worm onto the fixative so it is being fixed from below at the same time it is being fixed from above. Immediately before introducing the worms to the fixing container add a little ice-cold fixative or seawater to just melt the surface. Now float or coax the flatworm onto a piece of filter paper (or bond paper) that is porous yet strong enough to hold together when wet. Place the paper with the animal directly onto the flat, frozen surface of the fixative. The animal will flatten down, but may also ruffle or hump as the fixative makes contact. Use a paintbrush or glass slide to keep the animal flat for the few seconds it takes for it to be fixed. Without further disturbing the worm allow the fixative to thaw and leave them in this fixative for at least 24 hours. For longer-term preservation and storage worms may then be transferred to 70% ethanol. This method ensures that animals do not disintegrate and are fixed flat for microscopical examination.

A modern addition

These days researchers like to have samples of the animal’s DNA for molecular analyses, so another specimen should be fixed directly in 95% or 100% ethanol (or just cut a small piece off the posterior end and let the worm go).