Crayfish Collection for NSW

Materials and Methods

Freshwater crayfish were collected in NSW from 2005 to 2008 for the field guide. Only newly collected crayfish were used, no effort was made to view previous collections. As much of NSW as physically possible was sampled but collection was concentrated from the western slopes of the great divide to the sea. Unfortunately there was no set pattern for our collection activities so vast areas still remain unsampled. However collection and sampling will continue for the next 4 or more years. Our base plan on each expedition was to head for a known crayfish area by a new unsampled area and return by a new unsampled area. We would stop at every likely site along the way and collect samples. Only the samples required were retained the rest were returned after examination. Generally samples were collected by hand, this involved turning rocks digging up burrows or spotlighting at night. For larger spiny crayfish in larger rivers meat on string or traps were utilised (see capture methods). Regardless of the capture method for the majority of the samples at least one specimen of every species was dug from its burrow to obtain burrow information.

At each capture site, a GPS position was recorded as was any road or river locational information. Each collection site was photographed and where possible burrow entrances were also photographed. Each specimen was individually packed on site, labelled with permanent marker and boxed for return alive to the laboratory. Additionally, site notes were recorded at each site as to burrow information, crayfish activaty info and general comments. A collection record was completed for each sample location (click here).

Laboratory

All specimens were returned alive to our office/laboratory at Port Stephens NSW. Each specimen was housed in glass aquaria and observed as to activates and actions. Observation of crayfish in aquaria is quite enlightening and valuable information was gained this way. Each specimen was photographed alive in the aquaria using 2 cameras. A Canon 400D digital, 10.5 megapixal fitted with an EF100 mm macro lens and a Pentax Optico mx fitted with a macro lens were used for all photography. Freshwater crayfish are not the easiest animals to photograph but using glass aquaria designed and set up for this task made life easier.

After a time from 3 days to 6 months crayfish were removed from aquaria and processed. Crayfish were weighed on digital scales, measured with digital vernier calipers and a DNA sample was taken. Crayfish were then euthanised by freezing and then photographed dead. The final photography was based around photographing the external characteristics of each specimen. These morphological characteristics are what distinguish the different species.

Crayfish were then vouchered in 75 – 85% ethanol and numbered and labelled. Specimens were then described. This involves examining and recording all the crayfish’s external characteristics and comparing those characteristics with my lists for known species. There is variation between the same species in the same region and from those of other regions so this is not an exact science but an initial assumptions can be confirmed with DNA results.

After vouchering and labelling of specimens the specimens are described. A male representative sample of all specimens is removed from the preservative and examined under a microscope. External morphological characteristics are examined and recorded.

Once all these characteristics have been recorded all other specimens from that collection site are examined to confirm/alter data compiled.

This information is then compared with the known descriptions of known species. If they correspond then confirmation is awaited from the DNA sequencing. If the description does not correspond with any previously described species then further study is required.

This will involve more detailed examination of all specimens and further collection of samples to ascertain distribution and variation of morphology across distribution, etc.

DNA testing

All specimens collected are DNA tested. The test results are used for a vast range of purposes. Firstly we have identification which is critical, we also have relationships which can also be seen with DNA results. As part of the Australian Crayfish Project every species of crayfish in Australia will be collected and DNA tested. This will add to existing data bases and create a full DNA data base for all Australian species. Not just every species is being tested, we are also testing the same species from different areas. For example the same species from different rivers, catchments, altitudes or habitats, etc.

All DNA testing for this project is being carried out by the Carnegie Museum of Natural History, Pittsburgh USA. We work with Dr James (Jim) Fetzner the Assistant Curator of Crustacea, Biodiversity Services Facility, Section of Invertebrate Zoology, Carnegie Museum of Natural History USA. We are extremely lucky to have the support and assistance of the Museum in processing the thousands of DNA samples required by this massive project.

All DNA testing requires a sample from each crayfish to be taken and preserved in DNA tubes. These tubes are small ... roughly 50 mm high by 10 mm diameter and half filled with cell lysis buffer. This is a clear liquid used to preserve the tissue samples. Once filled with a sample these tubes can be kept for about 3 months at room temperature and then sent to the laboratory for DNA testing. They come in a box of 81 and it will generally take us several months to collect samples so we just send off the samples in full box loads every few months.

Field Collection Protocol for Crayfish DNA Samples

Samples should only be taken from live or freshly dissected individuals. Typically, samples that have been sitting (dead) at room temperature (or higher) for even moderately short periods (an hour or more) will tend to yield little DNA because the DNA will degrade very rapidly in the absence of some form of preservation. Therefore, it is crucial that DNA samples be taken as soon as possible from dead dying, or fresh organisms and placed directly into Cell Lysis Buffer (CLB) after appropriately cutting the tissue (see below). Muscle tissue seems to work best. Abdominal, leg, or chela muscle can all be used. Tissue needs to be taken before placing specimens in ethanol.
Tissue samples should be minced or broken up before placing them into the CLB. This allows the buffer to infuse into the tissues, which helps preserve the DNA. If the tissues are placed into the buffer in one large mass, it will most likely rot (which is extremely bad for DNA), especially when you are collecting in the field for extended periods and can not get to the DNA extraction steps for a while. The CLB contains a high concentration of EDTA (a chelator) that binds the free metal ions (i.e., Mg*², Ca*², etc.) in solution. DNases (the nasty enzymes that destroy the precious DNA) need these ions in order to function properly, so removing them preserves the DNA.
Typically you want to add only a small amount of tissue to the CLB buffer. Adding too much tissue to the tube (a common mistake people make) can cause degradation of the DNA and rotting of the tissues. Tubes have been supplied with 900 ul of CLB, typically you will want to add a maximum of 0.5 to 1.0 cm³ cube of tissue (minced up of course) to this volume of CLB. Any more than this can cause problems. If you have extra tissue it is always best to use another tube (not good if tubes are limited) or just trash the extra tissue (best), rather than trying to stuff it all into the same tube.
After adding the tissue to the CLB, you should invert the tubes rabidly several times to mix the tissue and buffer well. Make sure that all of the tissue is floating in the CLB and not stuck to the sides or top of the tubes. Occasional mixing for the first few hours after adding the sample will aid in breaking up and dissolving the tissues. After a few hours you should notice that the clumps of tissue disappear (this is good), which means that the cells are lysing and the DNA is being released into the EDTA preservative. For cut up legs (my preferred option) you will notice that they become hollow. Once the tissues are in the CLB they can be stored at ambient temperatures for extended periods (weeks to months) and still yield high quality DNA. Even so, it is recommended finishing the extraction as soon as possible, so samples should be shipped to the lab ASAP after collection. The buffer itself (before adding the tissue) has a shelf life of up to one year.
Be sure to label the tubes so that accurate records can be kept for each sample. Typically you should label the top of the cap and the sides of the tube as a backup measure in case one label gets wiped off for some reason. Typically, a simple number will suffice on the tubes. Theses numbers are then cross referenced with your field collection notes and databases.
Vouchered specimens are also important. This will allow at a later date verification that the DNA sequences are indeed from the correct species. It is essential to cross reference DNA sample and vouchered sample to ensure easy identification of which DNA can from which specimen.

Collecting Tissues from Freshwater Crayfish Legs

Using the legs of the freshwater crayfish as the source of tissue for DNA samples is the preferred method. Crayfish are exceptionally resilient, they have the ability to regenerate lost appendages. Those crayfish that a DNA sample/leg are taken from and returned to the wild will not be seriously compromised by the lose of a part or whole leg if you only remove the second walking leg.

Starting from the rear of the crayfish the first 2 sets of legs are walking legs, these four legs all end in a pointed segment. The first set of walking leg are essential for stability, bracing, reach and fast movement so, they should not be removed. If you do use these legs then the abilities of your crayfish will be reduced, this is not a major problem as the crayfish will adapt and compensate with its other legs. The second set of walking legs are support legs and not so essential so it is recommended to use these legs. It is one of these legs that should be utilised to reduce any adverse effect on the crayfish. The next 2 sets of legs are feeding legs with the small nipper on the end. These are essential for feeding and burrow construction so should be avoided.

Specimens that are being retained also benefit from removal of only that leg, retaining the structural integrity of the whole crayfish allows researchers at a future time to fully examine the crayfish without the loss of any major components and if the legs need to be examine there is an exact duplicate of the lost leg on the other side of the crayfish.