<commit_before>---
layout: single
title: "Field to Database: which info do we collect?"
date: 2017-06-05
type: post
published: false
categories: ["Tutorial"]
tags: ["Biodiversity"]
excerpt: "A write-up on data collection for large biodiversity surveys"
---


## Foreword

This is our current workflow, it has been modified and perfected over dozen of field expeditions in various levels of sophistication and equipment availability. With this setup and workflow a team of 5-10 people can collect and process 150 to 400 specimens/lots a day. It has mostly been developed by <NAME> but other people have contributed ideas and improvements.

This workflow changes regularly, this is a write-up of the current one.

## What are we trying to accomplish?

When we are in the field to collect animals, 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 damanged by fixation methods (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.

## The constraints

Time in the field is precious. We want to work as efficiently as possible to maximize the number of specimens we capture. The process needs to be as streamilined as possible to capture as much information as possible in little time.

## 0. Before going to the field



## 1. The collection

### Rule #1: 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 sink, this also applies to their lids. Using glass jars with metal lids (e.g., jam jars) work best. If diving/snorkelig do not forget to fill them with water at the surface. If collecting in the intertidal, do not fill the jars with water to the top as they 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. Ziplock bags are useful, but they float. The ziplock bags with a zipper are not good, as inevitably the zipper will 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 would 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 put in jars with.
* Crabs tend to fight with each others, and lose arms and legs in the
  process.
* Several worm phylum produce also a lot of mucuous (nemeretean) that
  can be problematic to other specimens.
* Sponges have lots of secondary compounds that will kill other organisms. Keep
  them separate.

If coming back from a dive, stabalize what you collect. 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, in the shade to avoid the water to get too warm. If in really hot climate and long boat rides, having a little ice in the cooler can be useful. Otherwise, putting sea water in the cooler would be enough to avoid the temperature to rise too much during the trip back to the lab.

### Rule #2: 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 stabalizing 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 record the information.


As soon as possible, after stabilizing the animals just after the dive, write down on you slate any notes that will help you remember where you found the animals you collected. Writing down about the environment can also be useful (coral cover, species of coral or algae making up the bottom, etc.)


## 2. Stabalizing at the lab

### How to set up the lab?

At least 3 zones:

- wet and messy
- photo station
- relaxation station
- tissue subsampling station
- data entry


### The tools needed in the lab

* Sink
* Mops
* spoons
* twizers
* 5 gallon buckets
* 6-packs
* plastic pipettes
* paint brushes
* toothbrush
* white board/easel

* fridge/freezer

* 2mL vials

### Chemicals needed

* Magnesium chloride
* Clove oil
* Menthol
* Ethanol 75% and 95%
* Formalin

### Rule #3: Keep your animals alive

Once you arrive at the lab, 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 transfering your specimens from the field into the containers in the lab, be mindful of the thermic 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.

As soon as the specimens are transfered 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. You may want to 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 halimeda, before picking a dead coral head, that could represent 3 stations. The GPS coordinates would be the same, but the depths and habitat descriptions would be different.

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.

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 tissue sample, get preserved) 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.


### Rule #4: Assign field numbers early

It is good practice to assign field numbers early, so it is easier to keep track of the specimen as it goes through processing.

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. It should be unique for the institution. We currently use a 3 or 4 letter prefix folowed by a 2-digit number representing the year. For instance, last year we went to Bocas del Toro and used the prefix "BCS16-".

At this stage, we assign a field number to each specimen. For the medium size animals (2-10 cm), we usually keep them in the sandwich boxes. For the smaller ones, (2-20 mm), we use the "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. Except for some unusual circumstances, the answer will be "yes" to
  all of these questions.
* any notes about the specimen that are important. 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...

### Rule #5: 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.

The specimen should be photographed individually. The specimens should always be in submersed in water when photograhed. 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 them should include the full specimen. Additional photos can be taken to focus on some morhplogical detail. Different views might be needed (dorsal/ventral/lateral) ddepending 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 taking pictures of the animal alive and relaxed may be needed.


### Rule #6: Relax the specimens properly

We typically use 3 relaxants for marine invertebrates: magnesium chloride, clove oil, and menthol. Additonally, 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, disolve 75g of 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 individuals. Magnesium chloride can be used on molluscs, annelids and other worms (except plathyhelminthes), echinoderms.

Clove oil is used on arthropods. The clove oil from leaf extract is cheaper than from buds but does not work. A couple of drops in 200mL is all what is needd. Most shrimps react very quickly (but there are exceptions), while for crabs, and pagurids it can vary quickly from a few mintues to several hours. Intertidal species usually take the longest. It's generally common to anesthesize arthropods before taking their pictures, however, it is really important to rinse and gently brush them to remove as much oil as possible before transfering them to the photo tank. The oily residues show up on the photos and can't be removed. For anesthesized 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 disolves plastic. If you relax arthropods into plastic cups, change them regularly. It also disolves and smears the writing on the plastic labels we use. Be very careful to not spill or soack the labels into the clove oil solution.

Menthol crystals do not disolve well in water. It's more practical to disolve a a few crystals into ethanol, and add a few drops of this solution into sea water. It will recrystalize at the surface of the water but enough will go in solution. Use menthol to relax sea anemones and other cnidaries, and tunicates.

### Rule #7: Get tissue samples for genetic analysis

Getting tissue samples for genetic analysis is an important step of the 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 tools and the general 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 allow 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 (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 good 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 feets for sea stars and sea urchins.
* Arthrpods: for shrimps it's best to take pleopods, but take care of keeping
  one side intact. For (larger) crabs, take meat from the articulations between
  the leg and the carapace.

If preserving tissues for RNA additional precautionary measures need to be taken. For instance, magnesium catalyzes RNA lysis, therefore the 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 are sued to working with it for advice.

### Rule #8: 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 appendadages can fall off a few months after being collected. Using lower percentages doesn't guarantee proper fixation.

Specimens should be fixed in about 3 to 5 times the (soft-tissue) volume of the specimen.

We use whirl-pak 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 that with spines or other sharp protusions, 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 can really help 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 twizzers, pry the valves open.

Soft bodied organisms including worms, nudibranchs, cnidarians, ctenophores, tunicates, do not get preserved in ethanol but in formalin. We routlinely 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 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.

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.


### Rule #9: Bring back the specimens safely


## 3. Misc

### Rule # Keep the lab clean and organized

Clean the containers (glass jars from the field), plastic tubs from the lab all the time. Rince them with fresh water and 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

* take pictures of your field data sheets
* take pictures of your station data


### Update the workflow to fit the space you have available
<commit_msg>add paragraph about data sheets and labels<commit_after>---
layout: single
title: "Field to Database: which info do we collect?"
date: 2017-06-05
type: post
published: false
categories: ["Tutorial"]
tags: ["Biodiversity"]
excerpt: "A write-up on data collection for large biodiversity surveys"
---


## Foreword

This is our current workflow, it has been modified and perfected over dozen of field expeditions in various levels of sophistication and equipment availability. With this setup and workflow a team of 5-10 people can collect and process 150 to 400 specimens/lots a day. It has mostly been developed by <NAME> but other people have contributed ideas and improvements.

This workflow changes regularly, this is a write-up of the current one.

## What are we trying to accomplish?

When we are in the field to collect animals, 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 damanged by fixation methods (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.

## The constraints

Time in the field is precious. We want to work as efficiently as possible to maximize the number of specimens we capture. The process needs to be as streamilined as possible to capture as much information as possible in little time.

## 0. Before going to the field

### Print the datasheets and the labels

To enter data in the field, we use pre-printed datasheets. We 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 also capture who is filling up the sheet.

To go with these sheets, we also print field labels. We print the field labels in multiples of the number of lines there are in the sheet, and they are clipped together. These can then 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 never write them
  by hand (unless we run out), and so the risk of having duplicate numbers is
  very low.
- everyone uses the same system. It's a very easy to learn system, that even
  people who are not familiar with who participate in our expedition can learn
  quickly.
- the specimens are tracked 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 small numbers makes it easier to communicate with the
  rest of the group ("Are you sure BCS16-0126 is Alpheus pinkfloydi?" is easier
  than "Are you sure 3214546431432153424325412365245 is Alpheus pinkfloydi?"")

## 1. The collection

### Rule #1: 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 sink, this also applies to their lids. Using glass jars with metal lids (e.g., jam jars) work best. If diving/snorkelig do not forget to fill them with water at the surface. If collecting in the intertidal, do not fill the jars with water to the top as they 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. Ziplock bags are useful, but they float. The ziplock bags with a zipper are not good, as inevitably the zipper will 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 would 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 put in jars with.
* Crabs tend to fight with each others, and lose arms and legs in the
  process.
* Several worm phylum produce also a lot of mucuous (nemeretean) that
  can be problematic to other specimens.
* Sponges have lots of secondary compounds that will kill other organisms. Keep
  them separate.

If coming back from a dive, stabalize what you collect. 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, in the shade to avoid the water to get too warm. If in really hot climate and long boat rides, having a little ice in the cooler can be useful. Otherwise, putting sea water in the cooler would be enough to avoid the temperature to rise too much during the trip back to the lab.

### Rule #2: 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 stabalizing 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 record the information.


As soon as possible, after stabilizing the animals just after the dive, write down on you slate any notes that will help you remember where you found the animals you collected. Writing down about the environment can also be useful (coral cover, species of coral or algae making up the bottom, etc.)


## 2. Stabalizing at the lab

### How to set up the lab?

At least 3 zones:

- wet and messy
- photo station
- relaxation station
- tissue subsampling station
- data entry


### The tools needed in the lab

* Sink
* Mops
* spoons
* twizers
* 5 gallon buckets
* 6-packs
* plastic pipettes
* paint brushes
* toothbrush
* white board/easel

* fridge/freezer

* 2mL vials

### Chemicals needed

* Magnesium chloride
* Clove oil
* Menthol
* Ethanol 75% and 95%
* Formalin

### Rule #3: Keep your animals alive

Once you arrive at the lab, 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 transfering your specimens from the field into the containers in the lab, be mindful of the thermic 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.

As soon as the specimens are transfered 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. You may want to 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 halimeda, before picking a dead coral head, that could represent 3 stations. The GPS coordinates would be the same, but the depths and habitat descriptions would be different.

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.

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 tissue sample, get preserved) 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.


### Rule #4: Assign field numbers early

It is good practice to assign field numbers early, so it is easier to keep track of the specimen as it goes through processing.

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. It should be unique for the institution. We currently use a 3 or 4 letter prefix folowed by a 2-digit number representing the year. For instance, last year we went to Bocas del Toro and used the prefix "BCS16-".

At this stage, we assign a field number to each specimen. For the medium size animals (2-10 cm), we usually keep them in the sandwich boxes. For the smaller ones, (2-20 mm), we use the "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. Except for some unusual circumstances, the answer will be "yes" to
  all of these questions.
* any notes about the specimen that are important. 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...

### Rule #5: 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.

The specimen should be photographed individually. The specimens should always be in submersed in water when photograhed. 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 them should include the full specimen. Additional photos can be taken to focus on some morhplogical detail. Different views might be needed (dorsal/ventral/lateral) ddepending 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 taking pictures of the animal alive and relaxed may be needed.


### Rule #6: Relax the specimens properly

We typically use 3 relaxants for marine invertebrates: magnesium chloride, clove oil, and menthol. Additonally, 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, disolve 75g of 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 individuals. Magnesium chloride can be used on molluscs, annelids and other worms (except plathyhelminthes), echinoderms.

Clove oil is used on arthropods. The clove oil from leaf extract is cheaper than from buds but does not work. A couple of drops in 200mL is all what is needd. Most shrimps react very quickly (but there are exceptions), while for crabs, and pagurids it can vary quickly from a few mintues to several hours. Intertidal species usually take the longest. It's generally common to anesthesize arthropods before taking their pictures, however, it is really important to rinse and gently brush them to remove as much oil as possible before transfering them to the photo tank. The oily residues show up on the photos and can't be removed. For anesthesized 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 disolves plastic. If you relax arthropods into plastic cups, change them regularly. It also disolves and smears the writing on the plastic labels we use. Be very careful to not spill or soack the labels into the clove oil solution.

Menthol crystals do not disolve well in water. It's more practical to disolve a a few crystals into ethanol, and add a few drops of this solution into sea water. It will recrystalize at the surface of the water but enough will go in solution. Use menthol to relax sea anemones and other cnidaries, and tunicates.

### Rule #7: Get tissue samples for genetic analysis

Getting tissue samples for genetic analysis is an important step of the 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 tools and the general 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 allow 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 (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 good 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 feets for sea stars and sea urchins.
* Arthrpods: for shrimps it's best to take pleopods, but take care of keeping
  one side intact. For (larger) crabs, take meat from the articulations between
  the leg and the carapace.

If preserving tissues for RNA additional precautionary measures need to be taken. For instance, magnesium catalyzes RNA lysis, therefore the 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 are sued to working with it for advice.

### Rule #8: 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 appendadages can fall off a few months after being collected. Using lower percentages doesn't guarantee proper fixation.

Specimens should be fixed in about 3 to 5 times the (soft-tissue) volume of the specimen.

We use whirl-pak 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 that with spines or other sharp protusions, 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 can really help 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 twizzers, pry the valves open.

Soft bodied organisms including worms, nudibranchs, cnidarians, ctenophores, tunicates, do not get preserved in ethanol but in formalin. We routlinely 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 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.

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.


### Rule #9: Bring back the specimens safely


## 3. Misc

### Rule # Keep the lab clean and organized

Clean the containers (glass jars from the field), plastic tubs from the lab all the time. Rince them with fresh water and 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

* take pictures of your field data sheets
* take pictures of your station data


### Update the workflow to fit the space you have available

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