species listspongescnidarianspolychaetesmolluscscrustaceansbryozoansascidiansspecimen handling


The following section provides both general and taxa-specific specimen handling techniques for invertebrates likely to be found on artificial substrates such as docks and pilings.

General Techniques
· All references to formalin below mean formalin stock diluted 1:9 with seawater
· Mix alcohol with de-ionized water to avoid percipitates
· Always completely submerge specimens in preservative and make sure the specimen is not too large for the jar
· Preserving solutions (both formalin and alcohol) used to fix material rapidly becomes very acidic; if material cannot be processed promptly upon return from the field, it is advisable to change the preserving solution to avoid acidity problems; no material should remain in its initial fixing solution for more than one month
· Sort specimens and groups according to fixing requirements; do not mix hard and soft animals; some fragile specimens may be damaged or destroyed
· Soft-bodied animals or unique specimens should be sorted directly into individual specimen jars
· When labeling specimens during field collection, be aware that some live animals will eat or otherwise destroy paper labels
· Any material that may be required for DNA analysis must be either frozen or fixed in 100% ethanol
· When freezing to relax or store specimens, do not thaw and re-freeze them; defrost one, photograph, if necessary; then fix in preservative
· It is important to cross-reference any photographs to the actual specimen photographed; make sure that field labels record this
· Material which has been fixed properly in formalin can be transported damp without liquid, if it is sealed in containers

Labeling is an important step in collecting any specimen. Labels should include the collectors name, date, exact location, habitat type (e.g., wooden pier piling, floating dock, patch reef, etc.), and depth of collection. Detailed notes regarding the living color of the animal are essential for the positive identification of many invertebrate groups, as most animals lose all color upon preservation. Photographing the living specimen before preservation is ideal. Labels should be placed inside collection bags or bottles as soon as possible, preferably at the time of collection. Specimens should be placed on ice in the field or quickly transported in sea water to a laboratory for sorting and preservation. In most cases, specimens should be narcotized and preserved within eight hours.

Preservation Methods for Specific Taxa

If possible, photograph live specimens in situ to record colors and growth form. Some specimens will disintegrate when handled. Preserve in 100% alcohol, or freeze, then preserve.

Sea Anemones
If possible, photograph and relax in the field live specimens before fixing. Put in a jar with enough seawater to allow the specimens to fully expand, then freeze or add menthol or magnesium chloride (epsom salt) and leave overnight; fix in formalin by adding the correct amount to the frozen specimens making sure it mixes as it defrosts.

If possible, photograph live animals; narcotize large hydroids in menthol or magnesium chloride overnight prior to fixation. Fix in formalin; store in formalin or 70% alcohol.

Soft corals
If possible, photograph and relax specimens before fixing. Put in a jar with enough seawater to allow the specimens or expand fully, then freeze or adds menthol or magnesium chloride. Leave until relaxed, fix in formalin for a maximum of 12 hours; rinse thoroughly in water, store in 70% alcohol.

Flat worms [Platyhelminths]
If possible, specimens should be photographed alive. It is important that they are preserved as flat as possible. Specimens can be relaxed using menthol or magnesium chloride overnight; but this is not always successful; specimens often disintegrate. The best method is to freeze a small amount of formalin a jar; then place the specimen on top when it will freeze onto the surface of the formalin, die flat and be fixed at the same time. Add an appropriate amount of seawater to make up the solution.

Polychaete worms
These specimens can be fixed directly in formalin. Some larger species may need to be relaxed using menthol or magnesium chloride prior to fixing. Try to remove tube-dwelling species from their tubes to allow proper fixation; always retain the tubes. Many species will fragment; all fragments should be retained. Fix in formalin and store in formalin or 70% alcohol. In the case of species with calcareous tubes, transfer from formal to 70% alcohol within 24 hours of fixing.

If possible, photograph alive as living colors can be useful identification features. Fix hard species in formalin, if possible, then dry; store dried. Soft or lightly calcified species should be fixed in formalin (not more than a few days); store in 70% alcohol.

General: Most mollusks can simply be put directly into formalin to fix and are usually stored in formalin. It can be helpful to relax snails (gastropods) specimens.

Opisthobranchs (and other reduced-shell gastropods): Specimens must be photographed alive as form and color patterns are very important diagnostic features. Specimens must be relaxed before fixing. The best method for relaxing is to put specimen in a jar with enough seawater for it to crawl around with rhinophores, gills, etc. fully extended; freeze overnight. Add enough stock formalin to frozen seawater to make up solution of appropriate strength, make sure it is mixed as the seawater thaws. If freezing is impractical, use menthol, magnesium chloride in seawater, or iced seawater, overnight, to relax specimens. Fix in formalin; do not leave specimens in formalin for more than one week; store in 70% alcohol.

Bivalves: For species with shells that seal tightly, place a match stick or similar object between valves prior to fixation to insure that fixative reaches internal tissues. To get bivalves to gape, either warm until they relax enough or freeze them. Fix in formalin, store in formalin (except for species with very thin shells).

If possible, photograph living specimens, particularly shrimps. Specimens are best fixed alive. Remove hermit crabs from their shells and tube-dwelling species from the tubes prior to fixing. Avoid putting specimens with large claws in with other animals as they may grab and damage more fragile species. It is sometimes preferable to kill large crabs individually and them put them in a communal container to fix. Fix in formalin and store in formalin or 70% alcohol. Do not freeze crustaceans unless there is no other option, as they do not fix as well after having been frozen.

Echinoderms are not usually among introduced species and rarely found on docks and pilings. At any rate they should be photographed while alive. Asteroids, ophiuroids, and echinoids should be fixed in formalin and most can be dried; otherwise they should be stored in 70% alcohol. Sea cucumbers can be fixed in 100% alcohol and stored in 70% alcohol.

Tunicates [Ascidians]
Compound, colonial, or other gelatinous ascidians should be photographed alive as form and color patterns are very important diagnostic features. Large solitary ascidians should be relaxed before fixing; menthol or magnesium chloride in seawater overnight is usually effective. Large solitary ascidians may also need to have preservatives injected into them to insure adequate fixation; fix in formalin; store in 70% alcohol.

[Specimen handling information modified from: Hewitt, C.L. and R.B Martin. 2001. Revised protocols for baseline port surveys for introduced marine species - Survey design, sampling protocols, and specimen handling. Centre for Research on Introduced Marine Pests, Hobart, Tasmania. Technical Report 22.]


© 2002 Hawaii Biological Survey, Bishop Museum