Ichthyology 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
• Randy Singer - The Florida Museum of Natural History
• Kevin Swagel - The Field Museum of Natural History, Assistant Collections Manager - Fishes
• H.J. Walker - Scripps Institution of Oceanography, Ichthyology Collection Manager (Retired)

Fluid-Preservation Techniques

Preservation of tissue samples

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

• Tissues to sample: whole organism, fin clip (right side of organism), 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

• 10% buffered formalin (= 3.7% formaldehyde) (dilute with distilled, de-ionized water)

Buffers:

• sea water and marble chips, Borax, baking soda
• 4.0 g monobasic sodium phosphate monohydrate and 6.5 g of dibasic sodium phosphate anhydrate per 1 liter of solution consisting of one part commercial formaldehyde (37%) and nine parts of distilled or deionized water (Simmons, 2015)

• If keeping specimens in 10% formalin for long term storage, you’re done. However, be sure to check formalin pH routinely (??) to ensure no acidification is occurring.

Specimens that are always stored in formalin for long-term storage: eggs Specimens that are sometimes stored in formalin for long-term storage: larval fishes/amphibians

• If transferring to ethanol or isopropanol for long term storage, then the amount of time depends on size of specimens.

Some general recommendations based on experience:

• Small stream fishes (e.g., minnows, <10 cm), fix for a week or so.
• Medium-sized (e.g.,sturgeon, <70 cm), fix up to a month.
• Large (e.g., White Shark, ~3m), fix up to a couple months.

Note: A well-fixed specimen is firm to the touch.

• Inject with 10%-20% formalin for “larger” specimens.

Cut open body cavity for “very large” specimens (right side). If you take it out too soon - the specimen will rot. If you take it out too late - specimen will de-calcify.

Ethanol Fixation

Although not ideal, another option for fixation is 95% or 70% ethanol – drop specimens directly into fluid, store. When transferring from 10% formalin to ethanol (70-75% concentration) or isopropanol (50%) for long term preservation: Ideally, step up from water (until no more formalin remains in fluid), 25% ethanol, 50% ethanol, 75% or 70% ethanol (with appropriate modifications for 50% isopropanol). Some collections modify this step up procedure (e.g., one step in 35% or directly into 70%).

Fixation and Preservation (Marine fish eggs and larvae)

Historically, Smith and Richardson (1977) has been followed, preserving fish early-life stages in 5% formalin. Typically the plankton sample is washed from the cod end into a quart or gallon jar and filled three-quarters full with seawater. Full strength formalin (37%) is then injected into the sample 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.