Collecting methods are too varied to cover in detail, but we will demonstrate a number of them during the field excursion. In nearshore environments in Florida much of the habitat consists of sand, silt or mud. In these areas most animals live buried, and need to be exposed by digging/sieving. Hard bottoms, like jetties, oyster reefs, or mangrove roots, offer home to other species. Seagrass beds hold many species that can be collected by a net run through the grass bed. Initial collecting for learning and documenting what species occur in an area is best done qualitatively, i.e. without worrying about documenting the abundance and local distribution of various species. Once the species are well-enough known, quantitative studies of communities can also be pursued.
It is useful to keep animals that mess each other up in separate containers after collecting. Some mollusks slime (that’s no big deal to other mollusks, but can really foul a crab), crabs rip, many sea slugs and flatworms emit toxins that kill other organisms confined in the same container. The two biggest enemies of collected animals in confined containers are lack of oxygen and heat. It is best to keep few organisms per container and make sure there is ample air space above - animals in a closed jar filled with water to the top will quickly use up all the oxygen and suffocate.
There are three main components to thoroughly documenting collected animals: photography, subsampling, and voucher preparation.
1) A collecting event at a certain place and time, usually from a specific habitat, is assigned a Station Number (ex: AMB-St-005). This number is recorded with all its requisite information on the back of a pre-printed field sheet (see attached “field sheet”) and possibly in the spreadsheet which is saved on the computer (see attached “spreadsheet”). The Station Number links to information about the collection site:
A. Location description (country, state, city, island, address, name of location, description of how to find site). EX: Djibouti, Moucha Islands, ridge off Maskali Island
B. Date of collection
C. Collector(s)
D. Latitude / Longitude
E. Depth (or altitude)
F. Habitat
2) When coming in from the field, animals from the same Station are initially kept together, a label with the station number is added to their container for tracking, and are then sorted to morphospecies. Each morphospecies gets a pre-printed label containing the Field Number (ex: FLAR-004). There are two copies of each Field Number on each label (one for specimen voucher, one for tissue sample for DNA) and they match the numbers on the printed field sheet.
3) As a specimen is sorted for processing its information is entered next to its assigned field number on the sheet. This information includes:
A. Station Number
B. Taxon
C. Fixative to be used.
D. Whether the specimen will be photoed (and any relevant notes, ex: in situ photo)
E. Whether the specimen will be subsampled.
F. Number of specimens associated with that field number (if more than 1)
G. Microhabitat
H. General notes
4) The specimen then moves on to the photo area along with both copies of its field number tag. The first photo of the specimen includes at least part of the animal and all of the field label. This enables easy subsequent matching of photos with specimens, and also provides a scale if the printed field number is of a standard size. The photos following the photo of the field number all go with that number.
5) After photography (or if the animal is not going to be photographed) the specimen and its pair of Field Labels moves on to the relaxation/subsample/fixation area. Animals are anesthetized according to their phylum and/or taxon (different relaxants work on different groups).
6) Once the animal is thoroughly relaxed a subsample is taken (if applicable) and placed in a small (2 mL) plastic, “DNA” vial filled with 95-100% ethanol. One copy of the Field Label goes into the small vial with the subsample. This allows us to match the subsample with the voucher back at FLMNH.
7) The animal (voucher), along with the remaining copy of the Field Label goes into the fixative appropriate for that group. If no subsample was taken, both copies of the Field Label will go into the vial with the voucher. This is a further check as to whether or not a subsample was taken in the field.
8) As the field sheets are filled, photos or copies of it are made for backup.
9) After field work the field sheets are transcribed into the spreadsheet provided.
Some animals are best photod in the field, others, especially small, mobile ones, are easiest to photograph in the lab. For lab photography use a small aquarium, with just enough water to cover animal. A black background (velvet, felt) placed under the aquarium gives an even background (we will discuss tricks of getting this especially even). The field label, and a scale, gray card, color card, are useful to photograph with the subject in the first photo. If the animal moves too rapidly to photograph it can be anesthetized or killed prior to photography.
Point and shoot cameras need to be set to macro mode. Flashes on these often give uneven lighting at close range, so if you use them try to get other lighting sources (natural light, strong LED, etc). SLR cameras with macro lenses are ideal for photographing small animals, especially when combined with flashes. If you shoot with SLR, synchronize the flashes, and set the f-stop around 16-20. Lower f-stops give less depth of field, while higher f-stops lead to low resolution, the 16-20 range is a sweet spot of compromise. We like to manually focus when photographing with SLR, as most auto-focus functions are not sufficiently responsive. Make sure you select the approriate white balance and high resolution, low compression.
Several anesthetics are commonly used with marine animals:
Magnesium chloride (MgCl2): prepared in fresh water at 7.5 weight % (75 gr of MgCl2 hexahydrate to 1 L water), so that it is isoosmotic with seawater. It is gradually added to seawater, typically for a final 50-50 concentration. Mg replacing Ca in muscles and nerves paralyzing the animal. MgCl2 works well with most marine invertebrates: worms, mollusks, anemones, echinoderms, etc.
Clove oil a few drops mixed up in sea water, then this murky solution added to a bowl with animals works great on crustaceans especially.
Menthol add to dish with animals by either sprinkling crushed crystals on top or adding drops of concentrated menthol solution prepared in ethanol. Menthol works especially well for cnidarians and ascidians. Even better, add menthol on top of dish of MgCl2 solution – works very well for such difficult to relax animals like sea cucumbers.
Cooling / freezing : Prevents autotomization in many and is a relatively humane way of killing animals, but will not commonly lead to expansion of contracted animals. Great for crabs.
The purpose of fixation is to fix tissues for long-term storage and study. Formalin or similar fixatives are necessary for quality fixation and for most groups where detailed anatomy or histology is desired, but formalin makes DNA extraction very difficult at best. Ethanol is fine as a fixative for groups where only external characters or gross anatomical features are used in taxonomy (e.g. most crustaceans and sponges), is more pleasant and less hazardous to work with, and DNA can be extracted from the specimen. For all specimens it is ideal to take a small tissue sample fixed separately for DNA extraction, but this is especially important for samples that get fixed in formalin.
When fixing large animal (such as a large sea cucumber, sponge, soft coral) in ethanol, it is best to use 95% ethanol to counteract the water content of the animal - you can eyeball. You should always use enough fixative - at least 3X volume of specimen, to make sure that the final concentration of fixative is adequate (70% for ethanol, 5-10% for formalin). For larger animals (>5 cm in all dimensions) it is important to make sure the fixative penetrates, by injecting or cutting them open.
Formalin: used generally at 5-10% strength of the industrial “formalin” solution (itself 38% formaldehyde gas dissolved in water). Thus 10% “formalin” is 3.8% formaldehyde. For marine life, mix formalin with sea water; for freshwater animals mix with freshwater. For any group with calcareous parts buffer formalin, as the liquid turns acidic (formic acid) with age. Buffering can be done with fine CaCO3 powder or a pinch of borax.
Ethanol: Bulk alcohol usually comes at 95% concentration. We use 75% ethanol for routine fixation, 95% for routine bulk genetic fixation, and 95-100% for fixing small genetic subsamples. Mix ethanol with distilled/deionized/soft water to dilute it. Methanol is not good for fixation; isopropyl is OK, but not optimal.
The following groups need formalin for proper fixation: algae (if fixed wet), anemones, worms, cephalopods, ascidians, fish.
The following groups should be fixed in alcohol: sponges, cnidarians other than anemones, mollusks, crustaceans, echinoderms
Subsampling is useful because you get better DNA preservation in small tissue pieces than in larger, whole specimens, and because it allows samples to be stored in freezers, where DNA quality remains high. A tissue sample should be <1-2 mm across to allow penetration of alcohol. Little tissue is needed to get DNA, but it is useful to have a couple of these pieces to allow repeated DNA extractions. To take a subsample, slice off a small piece of the animal (end of foot in a gastropod, tube foot from a sea star) with a scalpel or razer, and place it directly into 95-100% ethanol in a subsample vial. Makes sure there is 10X the volume of ethanol to fix the sample. With small animals or those that deteriorate rapidly (sponges, flatworms, etc) it is important to fix a subsample (or the sample) before, or quickly after death as DNA can deteriorate rapidly. Robust animals (e.g. fish, mollusks) however yield quality DNA many hours after death.Information about collecting