This is the current workflow of the Division of Invertebrate Zoology, Florida Museum of Natural History since about 2015. It has been modified and perfected over dozens of expeditions in diverse field conditions, and equipment availability. It can be adapted to work in an hotel room, a ship, or a well-equipped marine lab. With this workflow a team of 5-10 people can collect and process 150-400 specimens a day. This workflow gets modified regularly depending on tools and technologies.
What are we trying to accomplish?
When we are in the field to collect specimens, we are trying to document the species that live in the area. We want to preserve physical specimens as evidence of their presence, live appearance to preserve characters that will damaged by fixation methods (e.g., loss of coloration), and tissues for genetic analyses. We also want to capture data about where the specimen was found, and when possible what the environment looked like.
Time in the field is precious. We want to work as efficiently as possible to maximize the diversity of the specimens we capture. The processing of the specimen collected needs to be as streamlined as possible to capture as much information from as many species from as many different habitats as possible.
0. Before going to the field
Print the datasheets and the labels
To enter data in the field, we use pre-printed datasheets. These sheets allow us to capture the station number, our best guess for the identity of the specimen collected, whether there is a photo, a tissue sample, and a voucher for this specimen, and any notes we want to make about this specimen. We will go into more details about this below.
These sheets are accompanied of pre-printed field labels. We print the field labels in multiples of the number of lines there are in the sheet (there are 60 entries per data sheets, and 20 field labels per label sheet). They are clipped together to make a unit that can easily be handled by a person, and everyone can have their own data sheet and associated field numbers.
This system allows us to make sure:
- the field numbers are unique. They are printed, we don’t write them by hand (unless we run out), so there is no risk of having duplicated field numbers.
- the field numbers are small enough that they fit in 2mL DNA vials.
- everyone uses the same system, and it’s easy to use and learn.
- the specimens are tracked with this number from the start. As soon as a specimen gets a field number assigned, it will stay with it for the rest of its museum life. This is the number that is associated with the photos and the tissue samples. Ideally this number should be a global unique identifier, but this is impractical in the field. Having these field numbers makes it easier to communicate with the rest of the group (“Are you sure PKLS16-0126 is Synalpheus pinkfloydi?” is easier than “Are you sure 301fcb77-4e52-4817-94ce-91fd9d81a881 is Synalpheus pinkfloydi?”)
Packing and preparing the lab: the tools and equipment needed
Optional but makes life easier:
- Fridge and freezer
- Mops and other cleaning supplies
- 5-gallon buckets
- White board/easel and markers
- Trash cans
- Paper towels
- LED lights (we love this one)
- Plastic containers (different sizes)
- Plastic cups
- Tweezers soft, long
- Plastic pipettes
- Paint brushes
- 2mL plastic vials with screw caps
- Whirlpak bags: 4oz, 18oz and 42oz
- Permanent markers (ethanol resistant)
- India ink pens for labels and pencils
- Label paper
- Magnesium chloride hexahydrate
- Clove oil (from buds)
- Menthol crystals
- Ethanol 75% and 95%
How to set up the lab?
The organization of the lab depends of the space and equipment available. There are typically 5 areas, that should be arranged in a way that makes it easy for specimens to flow through each step, and that people can move around the lab comfortably and safely.
The sorting station
- sorting trays
- soft forceps
- long forceps
This is a wet and messy area. This is where bulk samples (rubble brought back from the field, clumps of algae) are picked through to find the small specimens. Having good lighting is important. Use forceps (soft) and spoons to transfer the specimens into clean water in 6-packs or other individual containers, assign them station numbers or field numbers right away to keep track of their origin.
The photo station
- small paint brushes and toothbrush
- spoons, plastic pipettes
- coffee filters and fennel or filtered sea water supply
- turkey baster
- good lights
- bucket to discard dirty water
- alcohol to clean the glass of the photo tank
This is a semi-wet area. This should be organized to avoid spills and keep the photo equipment as dry as possible. Wiping the camera with a lightly wet cloth, followed by a dry cloth, at the end of the day to rinse off salt significantly. increases the life of the equipment.
The relaxing station
This is a wet area, it should be close to the photo station, and usually the person in charge of the photo station is also responsible of supervising the relaxation: some specimens will need to be relaxed before photography, while the others will go to relaxation after getting their picture taken.
Make sure to label containers that contain magnesium chloride to separate them from the ones that only contain sea water. Magnesium chloride solution can be re-used for a couple of days, especially if the container get cleaned regularly.
Be careful clove oil doesn’t get in contact with field labels (see below).
Take care of keeping the specimens associated with their labels.
The tissue sampling station
- Scissors (small, like iris scissors)
- Molecular grade ethanol (95%)
This station should be after relaxation and before fixation. It is a semi-dry area. It needs to be kept clean and tidy. It is usually best if one person is in charge of this task.
The fixation station
- carboy of 70% ethanol
- carboy of 10% formalin
- label paper + India ink pen/pencil
- 5 gallon buckets or other large containers
The fixation station is where specimens go into ethanol or formalin. Safety precaution for handling these fluids should be followed. The top of the whirlpak should be folded enough times to ensure a proper seal. The correct fixative should be used for the specimen. Adding a label to formalin-fixed samples makes it easier to deal with these specimens back at the museum (see below). The whirlpak bags filled with fixative should be stored in a plastic bags, themselves placed in buckets or plastic bins, in order to contain possible leaks.
- A computer station a dry area of the lab where laptops can be used to check field guides and species descriptions to help with identifications, backup photos and data.
- A microscope station a dry area where small specimens can be looked at/sorted.
Keep the lab clean and organized
With the daily activities around the lab, it is important to keep the lab clean. It means making sure the floors get swept/mopped, the containers (glass jars from the field), plastic tubs from the lab should be rinsed and cleaned as soon as they are not in use. Rinse them with fresh water and air dry them. Do not use any soap, some of it gets trap in the glass/plastic and can be toxic for the animals.
Backup your data
It is crucial to backup the data generated in the field. The field data sheets, the white board with the station information should be photographed every day (if not more often). These photos as well as the specimen photos should be copied on multiple laptops and hard drives, and if the internet connection of the lab allows it put them in a Dropbox folder. The photos of the data sheets and the spreadsheet of the station data should also be emailed to people who stayed back home.
1. The collection
Things you will need
- glass jars with metal lids
- Ziploc bags
- underwater paper + pencil
Keep the specimens you are collecting alive and in good shape.
While in the field, underwater or on the intertidal, bring with you as many containers as possible. The ideal containers (and their lids) sink. Using glass jars with metal lids (e.g., jam jars) works best. If diving/snorkeling do not forget to fill them with water at the surface. If collecting on the intertidal, do not fill the jars with water to the top as the water will go anoxic quickly, killing whatever they contain. Be careful to not bang the collecting bag or to put hammers in your bag to avoid breaking the jars. Ziploc bags are useful, but they float (and they always end up with air bubbles trapped in them). The Ziploc bags with a slide-zipper are not good: the zipper will inevitably come off.
Keep animals separated as much as possible in your jars.
- Molluscs (especially snails) should be in their own jar. They tend to produce a lot of mucus that damage or asphyxiate other specimens.
- Many species of nudibranchs (especially phylidids) concentrate toxins of the sponges they feed on. They need to be on their own as these toxins will kill most species they will be in jars with.
- Crabs tend to fight with each others, and lose arms and legs in the process.
- Several worm phylum also produce a lot of mucous (nemeretean) that can be problematic to other specimens.
- Sponges have lots of secondary compounds that will kill other organisms. Keep them separate.
When coming back from a dive, stabilize what you collected. Split jars that have too many animals in it. Remove some water from the jars to allow for gas exchange and avoid anoxia. Place the jars in a cooler/ice chest, in the shade to avoid the water to get too warm. If in really hot climate and long boat rides, having ice in the cooler can be useful. Otherwise, putting sea water in the cooler can be enough to avoid the temperature to rise too much during the trip back to the lab.
Keep track of where you find what you collect
Usually, time underwater is very limited, so it is best to keep mental notes of where you found what you collected. However, diving with a slate for quick reminders is useful.
When stabilizing the specimen collected, add to the jar an identifier for the field site (e.g., A, B, C) if you are visiting multiple sites during a collecting trip, and/or a note to indicate who collected it. This will be helpful when you are back in the lab to sort who collected what, and record where the specimens were collected.
As soon as possible, after stabilizing the animals just after the dive, write down on your slate any notes that will help you remember where you found the animals you collected. Writing notes about the environment can also be useful (coral cover, species of coral or algae making up the bottom, etc.)
2. Stabilizing the specimens at the lab
Keep your animals alive
Once you arrive at the lab from the field, open up your jars, and transfer your specimens into plastic containers. Having a good assortment of sizes and shapes for containers is useful. Sandwich-size containers are great for most organisms: they provide a good surface for gas exchange, and are shallow enough that it’s easy to retrieve the specimens.
Plastic cups are good because it’s easy to get (even in remote locations), they are cheap, they don’t take a lot of room.
When transferring your specimens from the field into the containers in the lab, be mindful of the thermal shock. If the lab has A/C and the sea water from the lab is significantly cooler than the sea water you are bringing back from the field, the animals may not like the sudden change of water temperature. Mixing the sea waters to reduce the temperature difference may be necessary.
If you are dealing with very fragile animals that won’t stay alive very long or that are already dead, they need to be processed (get a field number assigned, get their photo taken, get a tissue sample, get them fixed) right away. Otherwise, at this point, things are relatively stable, and it’s a good time to rinse your diving gear/boat, take a shower, get a snack. You have many hours of work ahead of you. If the animals are put in clean containers, with fresh sea water, and the lab has A/C, most species can be kept in these containers for 12 hours or more.
Create station numbers
As soon as the specimens are transferred into their own containers, they should have a label with them that can be used to tell from which station/dive they are coming form and who collected them. This is a good time to create the station numbers that can be used to assign the origin of each specimen.
These station numbers should be displayed in the lab, so everyone can see them. They should have enough information to be identified unambiguously. Create as many station numbers as needed (they are cheap). For instance, if you did a dive where you spent the first 20 minutes at 15 m, and focused on going through coral rubble, and spent 60 minutes at 5 m going through an Halimeda patch, before picking a dead coral head, that could easily be split into 3 stations. The GPS coordinates would be the same, but the depths and habitat descriptions would be different.
At least once a day, the station numbers and associated information should be transferred into a spreadsheet. A check-mark or other indication should be added to the white board to indicate that the information listed on the board has been captured digitally. That way, if someone wants to add information about the site, they know it needs to be added to the spreadsheet.
Assign field numbers early
It is good practice to assign field numbers to specimens early to make it easier to keep track of them as they go through processing.
At this stage, we are usually splitting jars from the field, so it is crucial to make sure every animal has a label associated with it that indicates the (newly created) station number, and if needed who collected it. These labels can be done quickly with archival or underwater paper and pencil.
Currently, we use field labels printed on plastic paper. We print them ahead of time and bring them with us in the field. Each field number is duplicated, but one of them as the suffix “-dna”. The field numbers consist of a prefix that characterizes the field expedition. We currently use a 3 or 4 letter prefix followed by a 2-digit number representing the year. For instance, last year we went to Bocas del Toro and used the prefix “
Medium size animals (2-10 cm) are kept them in the sandwich boxes; while smaller ones, (2-20 mm), are kept in “6-packs”, each animal is placed in a cell with a number.
As soon as a specimen has a field number associated with it, we write down on the field data sheet:
- the station number where it came from
- our best guess for the identification
- whether a photograph of the specimen will be taken, whether a tissue sample will be taken, and whether we will keep the specimen as a voucher for the collection. In most cases, the answer will be “yes” to all of these questions.
- any relevant notes about the specimen. These may include: morphospecies match (if the species has already been collected during the expedition, it can be useful to refer to the field number to which it corresponds), the association (if it’s a parasite or a symbiont, we can put a note that indicates the field number of the partner of this biological association), the microhabitat (specific information about where the specimen was found), etc.
Take informative pictures of the specimens
The technical aspects of the setup and of the photo taking process will be the topic of a different post.
Specimens should be photographed individually and fully submersed in water. The first picture of the series should include the field label. This will be used to match the photos to the specimen later on. In most cases, we do not put a ruler on the photo because it considerably slows us down. Instead, we use the size of the print on the label to get a measurement that can be used to size the specimen. However, if we know that we found something particularly interesting (e.g., an undescribed species), we will add a scale in the photo.
Several photos for each specimen should be taken. Several of these photos should include the full specimen. Additional photos can be taken to focus on some morphological details. Different views might be needed (dorsal/ventral/lateral) depending on the organism. Pay attention to the lighting, and make sure that the photos are in focus.
Usually, it is best to take photos of the specimen when alive and exhibiting a “natural” pose. However, in some cases it is not possible and relaxing it before taking the photo may be necessary or preferred. For specimen of high importance, take pictures of the animal alive and relaxed.
Relax the specimens properly
We typically use 3 relaxants for marine invertebrates depending on the taxon: magnesium chloride, clove oil, and menthol. Additionally, for larger specimens, and arthropods in general, we also use the freezer/fridge.
Use magnesium chloride hexahydrate. The anhydrous form is too reactive for field conditions. For relaxation/anesthesia, dissolve 75g of hexahydrate magnesium chloride in 1L of fresh water, when available distilled/dionized water is best, but otherwise tap/rain water works. Use this solution in 1:1 mixture with sea water for anesthesia. The time it takes for organisms to respond to the magnesium chloride varies widely across taxon and even across individuals. Magnesium chloride can be used on molluscs, annelids and other worms (except plathyhelminthes), echinoderms.
Clove oil is used on arthropods. Clove oil from leaf extract is cheaper than from buds but does not work as well. A couple of drops in 200mL is all what is needed. Most shrimps react very quickly (but there are exceptions), while for crabs, and pagurids it can vary from a few minutes to several hours. Intertidal species usually take the longest. It’s generally common to anesthetize arthropods before taking their pictures, however, it is really important to rinse and gently brush them to remove as much oil as possible before transferring them to the photo tank. The oily residues show up on the photos and are a pain to clean when soiling the glass of the photo tank. For anesthetized arthropods, it is also important to pop-out the eyes of the crabs before taking their photos otherwise it doesn’t look good/natural. Clove oil dissolves plastic. If you relax arthropods into plastic cups, change them regularly. It also dissolves and smears the writing on the plastic labels we use. Be very careful to not spill or soak the labels into the clove oil solution.
Menthol crystals do not dissolve well in water. It’s more practical to dissolve a a few crystals into ethanol, and add a few drops of this solution into sea water. It will recrystallize at the surface of the water but enough will get into solution. Use menthol to relax cnidarians and tunicates.
Get tissue samples for genetic analysis
Getting tissue samples for genetic analysis is an important step of the biodiversity documentation process. The tissue sampled should be fresh, contain a good amount of material, be fixed properly to ensure high quality DNA, and taken in a way that is minimally destructive for the specimen.
The dissecting tools and the area where the subsampling takes place should be clean. The dissecting tools used for subsampling should be thoroughly cleaned between each specimen. If the safety conditions of the lab allow it, use a flame to sterilize the tools.
The piece of tissue taken should be about 1-3 mm in each dimension. It is best if this piece of tissue is fleshy (with as little skeletal elements as possible). Care should be taken to avoid epibionts and other associated organisms. These tissues will be used to get DNA material from the species in question not the species that live on it.
The tissue should be stored in high quality ethanol (ideally molecular grade). For most tissues, putting them directly in 95% works best. If you are dealing with thick/watery tissues, placing them in 75% for 24 hours before transferring to 95% is probably better.
With our double label system, it is at this time that we split the label, to put the half with the DNA suffix into the vial with the tissue.
When taking a piece of tissue on an animal, it needs to be done in a way that does not destroy morphological features that can be used for identification.
- Annelids (and other worms): both the head-end and the tail-end contain important features needed for species identification. Take a small section on one side of the animal. If the specimen is very small, use a few segments from the middle of the organism.
- Molluscs: usually a small sliver of the foot (for gastropods), mantel (for bivalves), or tip of a tentacle (for cephalopods) works well.
- Echinoderms: the tip of an arm for brittle stars and crinoids, one or a few tentacles for sea cucumbers (but make sure to not cut them right from the base, so it is possible to count the number of tentacles after they have been cutoff), tube feet for sea stars and sea urchins.
- Arthropods: for shrimps it’s best to take pleopods, but take care of keeping one side intact. For (larger) crabs, take muscle from the articulations between the legs and the carapace.
Tissue samples should be placed in freezer when available.
If preserving tissues for RNA additional care need to be taken. For instance, magnesium catalyzes RNA lysis, therefore specimens can’t be relaxed with magnesium chloride before preserving the tissues. In general, if you are preserving tissues for RNA analyses talk to people who have experienced doing it for advice.
Preserve the specimens properly
The way a specimen is preserved determines how easily it will be studied in the future, and even if it will be possible at all to identify it. Spending a few minutes to make sure the specimen is properly fixed can greatly improve the value of the specimens collected.
Using 70-80% ethanol is better. Higher percentages can make the specimens very stiff, for instance arthropods may not be flexible enough to be studied, or worse their appendages can fall off a few months after being collected.
Specimens should be fixed in about 3 to 5 times the (soft-tissue) volume of the specimen.
We use whirlpak bags which are very convenient, as they are resistant, relatively leak proof, and you can bring a lot of them without taking too much space in your luggage. However, specimens with spines or other sharp protrusions, should be wrapped in paper towels or fixed in different containers. If the specimen is large, it’s a good idea to tie the field label directly to it to avoid losing it. If the specimen has a high water content, changing the alcohol after 24 or 48 hours helps with fixation. Small and fragile specimens should be placed in plastic or glass vials of the appropriate size. Really fragile specimens can be held in place inside the jar with small amounts of paper towels/cotton/kimwipe.
Gastropods with thick operculums and bivalves can close up when placed in ethanol. Make sure that enough alcohol gets in contact with the tissues (hold the operculum open with tweezers, pry the valves open.
Soft bodied organisms including worms, nudibranchs, cnidarians, ctenophores, tunicates, do not get preserved in ethanol but in formalin. We routinely use 5-10% formalin solution (full strength formalin diluted in sea water). In addition of writing in the field notes, that the specimen is fixated in formalin, we also add a label (HCHO) with the specimen to indicate it is fixed in formalin. We also set aside the formalin-fixed specimens from the ethanol fixed ones. It really helps with the processing of the specimens when back at the museum. While most groups could potentially be fixed in formalin, given that it is hazardous, and that it makes the DNA practically unusable, we limit the taxa that get fixed in formalin. Some taxa shouldn’t get fixed in formalin as its acidity will destroy skeletal structures such as in echinoderms and bryozoans. Any specimen fixed in formalin should get a tissue sample taken.
Brittle stars should be fixed flat with all arms in the same orientation (comet shape) to facilitate future studies. Crinoids should be fixed with arms pushed down and flat. Sea cucumbers are best fixed with their body as straight as possible.
3. Back at the museum
Typically, we use 3 tables to keep track of the data from the field: a table for station data, a table for specimen data, and a table for photos.
The data from the data sheets need to be entered into spreadsheets. Most of the fields from the data sheets match the fields that are used in the collection database, making the transcription easy. The only field that need to be parsed is the “notes” that may need to be broken into several fields in the specimen table. Using a close correspondence between the fields used to enter data in the field and the data in the collection database ensures there is minimum manipulation to do to the data between the transcribed field data sheets and the database import.
Once the data is entered in the spreadsheet, the photos can be sorted, and matched to the specimens.
At this stage, before the data gets ingested into the database, we generally solicit the expertise of specialists to help with the identification of the material collected by sending them the photos of the specimens collected.