Herpetology Specimen Preparation

Statement of Purpose

These links and documents contain information about herpetology specimen preparation.

Contributors

Content generated during The American Society of Ichthyologists and Herpetologists (ASIH) Annual Joint Meeting - 2016, during an iDigBio sponsored workshop by the following individuals participating in the "Curation" working group of the aforementioned workshop:

• Sarah Huber - Virginia Institute of Marine Science, Ichthyology Collection Manager
• Justin Mann - Tulane University Biodiversity Research Institute, Ichthyology Collection Manager
• Katherine Pearson-Maslenikov - Burke Museum of Natural History, Ichthyology Collections Manager
• Susan Mochel - The Field Museum of Natural History, Assistant Collections Manager - Fishes
• Rob Robins - The Florida Museum of Natural History, Ichthyology Collection Manager
• Gregory Watkins-Colwell - Yale Peabody Museum, Herpetology/Ichthyology Collection Manager
• Kevin Swagel - The Field Museum of Natural History, Assistant Collections Manager - Fishes
• H.J. Walker - Scripps Institution of Oceanography, Ichthyology Collection Manager (Retired)

Fixation and Preservation

Preservation of tissue samples

Sub-sampling of specimens: DNA/RNA Preservation (must be done prior to fixation in formalin)

Tissues to sample (right side of organism): whole organism, fin clip, toe clip, muscle, liver, eggs) Link sample to specimen (see #Cataloging)

Storage of tissue samples:

• Dry (good for liquid nitrogen storage, -80 freezer)
• 95% ethanol/dry (pour off ethanol after 24 hours and freeze dry)
• 95% ethanol (pour off after 24-hours, re-fill with 95% prior to freezing)

Note: 100% ethanol can contain contaminates that may affect extraction RNA Later (then freeze)

• Tissue vials: cryovials, 1.8 or 2.0 ml Nalgene flat bottom and internal thread. Put a label in the vial, not just on the outside.

Parasites (project specific; may receive requests for these subsamples): Make notes of any external parasites, and keep everything together with sample.

Formalin Fixation

For amphibian larvae and eggs

Historically, Smith and Richardson (1977) has been followed, preserving fish early-life stages in 5% formalin. Full strength formalin (37%) is then injected into the sample container along with a saturated sodium borate solution as a buffer. For a liter (quart) 50 ml of full strength formalin is added along with 20 ml of saturated sodium borate solution. The container is then filled almost to the top with seawater and sealed. Two labels are generated for the jar, one outside and one inside, because the inside label will often be obscured by the plankton. The samples are then stored in a cool dark place, ideally around 65°F. Later, when the samples are sorted for early life history stages, the preservative can be changed to 3% buffered formalin for eggs and 70% ethanol for fish larvae. For DNA preservation, the formalin procedure should be skipped and the eggs and larvae should be put directly into ethanol.

For all others:

• Prepare a fixing tray which can be a shallow plastic container with a snap-on/air tight lid.
• Cover the bottom of the container with cheesecloth dampened with 10% buffered formalin.
• Injected buffered 10% formalin solution into body cavity and major muscles (avoid hitting nerves in legs).
• Spread specimen onto the cheesecloth and spread fingers and toes. The position in which the specimen is fixed will be permanent. Take care to ensure that all important characters can be seen.
  • For turtles, take care to extend the neck and limbs out of the shell in a natural position so that color patterns and scans can be examined.
  • For male lizards and snakes, evert the hemipenes by injecting formalin into the base of the tail and applying pressure. This technique requires practice.

""It is suggested that for at least some specimens of each taxon the mouth should be propped open to allow examination of the buccal cavity."" Because the position is permanent, keep in mind the final storage container. Lizard and salamander tails can be curved around the body in most cases. Whether snakes are coiled or fixed in elongated loops is a matter of personal and/or institutional preference as there are pros and cons to each (coils may fit in jars better, but loops may be easier to measure and for counting scales).

• Cover specimens with another layer of cheesecloth dampened with 10% buffered formalin. Additional fluid can be added to the container, but not more than 1 or 2 cm. The specimens should not be floating in fluid, but resting on cheesecloth. Specimens should rest in the fixing container for at least 2 days.

• After fixing, transfer specimens to a wash ethanol stage consisting of 50-70% ethanol for at least one week. Wash ethanol can be changed during this stage as needed. This stage permits formalin to diffuse from the specimen into solution.
• After washing is complete, transfer to 70% ethanol for long-term storage.