USGS


Observations on the Effects of Alcohol Vs. Formalin Storage
of Amphibian Larvae

Steve W. Gotte and Robert P. Reynolds

USGS Patuxent Wildlife Research Center
Division Of Amphibians And Reptiles
National Museum Of Natural History
Washington, D. C. 20560



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The following text represents an invited oral contribution presented in the workshop titled "Preservation and Curation of Early Life History Stages of Fishes, Amphibians, and Reptiles" at the meeting of the American Society of Ichthyologists and Herpetologists, held 26 June - 2 July 1997 in Seattle, Washington.

Because this is the only amphibian preservation talk at this workshop we include a quick overview of amphibian larval preservation and a brief history of larval collection management at the National Museum of Natural History as an example.

There are several general reasons for an increased interest in amphibian larvae in recent years. These include: a) increased awareness of the importance of biodiversity and the studies this has stimulated; b) increased work on amphibian ecology during the last couple of decades; and c) related to the previous two, increased research because of apparent amphibian population declines.

In addition, there are several specific reasons for increased interest in larval amphibians and resultant collection growth:

1) The larvae of many species have not been described; research to associate larvae with adults is being pursued by a number of researchers.

2) Larvae are often very difficult to identify, so voucher specimens must be collected in order to have properly identified synoptic collections available for comparison.

3) Even basic knowledge about geographic, ontogenetic, and environmental variation is lacking for most species. Work by Randy Jennings and Norm Scott (1993) and others has shown that the larval morphology of some species may be variable depending on developmental habitat.

4) Larvae are an 'accessible life history stage' for biodiversity studies and surveys. In some explosive breeders, such as spadefoots in the United States, and some canopy-dwelling tropical treefrogs, adults are only sampled easily for the few days they are active at the breeding sites, whereas the larvae are often accessible for weeks or months during their development.

History of Amphibian Larval Preservation at the United States National Museum (USNM)

As John Simmons has pointed out on a number of occasions (Simmons 1991, 1994), both the fixatives and preservatives used in museums have been determined as much by trial and error and tradition as by vigorous study. Before proceeding, let us clarify the difference between a fixative and a preservative.

According to R. W. Stoddart (1989), "The purpose of fixation is primarily to arrest the physical and chemical changes that would occur upon death of a tissue, and subsequently to such an extent as to preserve its gross form and appearance." Likewise, John Simmons (1995) stated, "Fixation is a chemical treatment of tissue that is used to prevent autolysis (the degradation of proteins into amino acids) and to coagulate cell contents into insoluble substances."

In contrast, "The preservative fluids used in spirit collections serve three main functions, which are to protect the specimen from decay or deterioration, to give it as 'normal' an appearance as possible and to afford it mechanical protection when handled" (Stoddart, 1989).

The following is a history of the larval fixation and preservation traditions at the USNM.

Table 1.

Years Fixative Preservative
<1890 ETOH/Spirits ETOH/Spirits
1890-1960 10% Formalin 70% ETOH
>1960 10% buffered Form. 10% buffered Form.

The first larvae in our collection are from the 1850s and were both fixed and preserved in ETOH (often in the form of distilled wine/brandy or rum).

Around the 1890s to the turn of the century, unbuffered formalin began to be used as a fixative. Specimens were then transferred to ETOH for preservation.

In his 1962 paper on preserving reptiles and amphibians, Bill Duellman stated that tadpoles should be put in 10% formalin, never alcohol. This is about the time that larvae were initially stored in 10% buffered formalin at the National Museum of Natural History.

Buffering the 10% formalin is essential to help ensure satisfactory long-term storage of amphibian larvae. We use a sodium phosphate monobasic/sodium phosphate dibasic (anhydrous) buffer (Quay, 1974) to maintain the pH of the 10% formalin near neutrality (pH 7.0). Formalin solutions that are either strongly acidic or too basic will have deleterious effects on the stored larvae: too acidic and larvae will become decalcified, whereas too basic and the specimens will clear. Current evidence suggests that the best solutions are neutral to slightly acidic; however, serious decalcification may start at a pH of around 6.5.

Fig1a.gif (39478 bytes) Figure 1. Photo of 1850s ETOH-preserved Rana catesbeiana in poor condition.
(click on image for larger picture)

This photo of bullfrog (Rana catesbeiana) tadpoles, which were collected in the 1850s and most likely both fixed and preserved in ETOH, shows that the specimens are extremely soft and mushy with few mouth parts remaining, and that the color is badly faded.  (Tying tags on individual larvae, as shown in this photo, is an old practice that is no longer used at the USNM. Currently, we lot catalogue larvae and place a single catalogue tag in the container with the specimen(s).)

Figure 2. Photo of ETOH-preserved
Hyla faber
(click on image for larger picture)
Fig2.gif (36894 bytes)

This photo of three ETOH-stored Hyla faber tadpoles collected on two different occasions by the same collector in Brazil in the 1920s shows the great variation possible in specimen quality after ca. 70 years in ETOH. While the two on the top are very soft and losing their interior structure and mouth parts, the one on the bottom is fairly firm and retains most of its interior structure and mouth parts.

Fig3.gif (36150 bytes) Figure 3. Photo of dehydrated Bufo marinus
(click on image for larger picture)

This photo of shriveled and deformed Bufo marinus tadpoles shows what most of our ETOH-stored tadpoles look like.

USNM Amphibian Larval Collection Today

The USNM formalin collection currently consists of ca. 1000 lots of salamander larvae and ca. 6000 lots of anuran larvae. We don't really know how many ETOH-stored larval-lots we have because larval specimens in the alcoholic anuran collection have not been inventoried. Our formalin collection experienced significant growth beginning in the early 1970s with accessions of larvae from Brazil and Panama as well as the eastern US. Since then, we have had major growth in larval collections from much of Latin America and the United States, and have added smaller collections from Asia, Africa, and Australia. Our larval collection has grown over 2500% in the last 25 years. Most impressive is that the collection has more than doubled in the last 5 years; i.e., we have added ca. 3500 lots in that time period. Much of this growth is attributable to the efforts of Roy McDiarmid and Ronn Altig and their associates to establish a comprehensive amphibian larval collection.

Larval Specimen Storage

Figure 4. Photo of two jars with vials. We no longer use the ground-glass jars with vials capped with Bakelite as shown on the left of the photo. The jar shown on the right depicts our current standard for formalin-stored larvae.
(click on image for larger picture)
Fig4a.gif (34827 bytes)

We use screw-top glass jars with polypropylene lids and polyethylene liners for storing our formalin larvae. When larvae are very small and/or delicate, or if a series of lots are related (such as a developmental series), we store them in glass shell vials with polyester fiber plugs, which are then submerged in formalin-filled screw top jars. Whether stored in a jar or a vial, we make sure there is an abundant volume of fluid relative to the volume of specimens. For birds and mammals, Quay (1974) suggested a specimen-to-formalin ratio by volume of 1:2 to 1:3. Because there is evidence that acidity increases as the specimen-to-formalin volume increases (Quay, 1974), we maintain a minimum amphibian larval specimen-to-formalin ratio by volume of 1:3.

We do not use Bakelite jar or vial lids. Besides the usual problems with Bakelite lids (cracking or becoming loose over time), Bakelite is a formaldehyde-based product and may react adversely with formalin preservatives. Likewise, bail-top jars with rubber gaskets are inappropriate and are not used as long-term larval storage containers because the gaskets soften and melt during extended exposure to formalin.

Alcohol Vs. Formalin Experiment

After our formalin collection was established and appropriate management procedures were developed, we decided to explore how best to deal with our large collection of larvae stored with adults in the ETOH collection. We had heard concerns that ETOH-stored larvae lost their mouthparts and generally disintegrated over time, and possibly it would be better to transfer them to formalin. However, we also received cautions that ETOH-stored larvae with intact keratinous mouthparts were prone to shed the mouthparts upon being changed to formalin and were otherwise damaged due to disruptions of the osmotic balance of the specimens.

We wanted to examine what affects changing preservatives had on larval amphibian specimens. Our experiment was in two stages: first we preserved formalin-fixed larvae in both 10% buffered formalin and 70% ETOH. This is the portion that we are reporting on at this workshop. The ultimate goal will be realized in the second stage, that is, to examine under controlled conditions the affects of transferring some of these ETOH-preserved specimens back to formalin, and to look for any adverse reactions that may result. This will establish baseline information that will be used to help us make decisions about either maintaining our ETOH-stored larvae in ETOH or transferring the ETOH-stored larvae in our collection to formalin.

We used three species of amphibian larvae for this study; the marbled salamander Ambystoma opacum (n=38), the bull frog Rana catesbeiana (n=25), and the green frog Rana clamitans (n=33).

Fig5.gif (33617 bytes) Figure 5. Photo of tadpoles, Rana clamitans (top) and R. catesbeiana (bottom).
(click on image for larger picture)

 

Figure 6. Photo of Ambystoma opacum larvae.
(click on image for larger picture)
Fig6.gif (29913 bytes)

We examined the larvae at the time of fixation and took several measurements (total length (mm), head length (mm), and head width (mm)) and qualitatively assessed general condition (texture, firmness), color patterns, and the condition of the mouthparts or gills. We repeated these same quantitative and qualitative measures on all specimens again after almost seven years. The non-parametric Mann-Whitney U and Wilcoxon Signed rank tests were used to statistically analyze the data, and significance was at the P=0.05 level.

Results

1) The mean measurements decreased within both of the treatments between the time of fixation and seven years later for all three species. That is, those larvae that were fixed and preserved in formalin shrank from the beginning to the end of the experiment, as did those that were fixed in formalin and preserved in ETOH. All differences were significant (P<0.05).

Table 2. Percent shrinkage of Rana larvae within treatments after seven years. The actual measurements had decreased significantly (p<0.05) for all larvae at the end of the experiment.

 

Total length

Body length

Head width

R. clamitans

 

 

 

Formalin

5.6%

3.7%

4.6%

ETOH

6.6%

5.9%

3.5%

 

 

 

 

R. catesbeiana

 

 

 

Formalin

6.3%

2.4%

4.5%

ETOH

5.1%

2.5%

2.3%

Table 3. Percent shrinkage of Ambystoma larvae within treatments after seven years. The actual measurements had decreased significantly (p<0.05) for all larvae at the end of the experiment.

 

Total length

Head width

A. opacum

 

 

Formalin

3.6%

11.0%

ETOH

5.2%

14.4%

2) At the end of seven years there was no significant difference (p>0.05) in the amount of shrinkage between specimens in the two preservatives in any of the three species.

3) Qualitative characters also seemed to show little difference between formalin-stored and ETOH-stored larvae. There was some yellowing of individual specimens within all treatments, but no obvious trends. Mouthparts seemed to be firm and well attached in all groups of tadpoles. Surprisingly, our ETOH-stored specimens did not show the shriveled "raisin" effects that many of the ETOH larvae in our collection show.

It is important to note that this experiment used large, robust species; results might not have been the same with smaller, more fragile species. Equally important is the fact that we used fresh, well-preserved specimens for these treatments. This preliminary short-term study on well-fixed specimens neither supports nor argues against the transfer of ETOH-stored larvae to 10% formalin. As previously stated, the second stage of this experiment is to transfer ETOH-stored larvae to 10% formalin to see if we can detect significant changes to the specimens. By doing this we hope to gain information which will aid us in making decisions regarding the best future storage medium for the larvae currently preserved in ETOH.

Formalin Larval Collection at the USNM - Overview of Current Procedures

Below is a brief review of current guidelines for the handling of newly collected amphibian larvae at the National Museum of Natural History:

Fix specimens in a relatively large volume of 10% buffered formalin in the field, transferring to smaller containers before packing for transport if needed. Transfer samples to a fresh solution of 10% buffered formalin when they arrive at the museum.

Maintain specimens in 10% buffered formalin as the preservative.

IT IS IMPORTANT IN BOTH FIXING AND STORING SPECIMENS THAT THERE BE AT LEAST A 1:3 SPECIMEN-TO-FORMALIN RATIO BY VOLUME. There is evidence that containers with high specimen-to-formalin ratios turn acidic faster. Check the pH in specimen containers periodically; the pH can change significantly during long-term storage, causing specimen damage.

In closing, the most important result revealed by these experiments so far is that it is imperative to fix the specimens well at the start; no matter what the preservation history, a well-fixed specimen will fare better than a poorly fixed one. Further, the finding that there were no discernable differences between the alcohol-and formalin-preserved specimens used in this study suggests that additional investigation should focus on determining the suitability of ETOH as a long-term storage medium for formalin-fixed anuran larvae.

Literature Cited

Duellman, W. E. 1962. Directions for preserving amphibians and reptiles. Pp. 37-40 In Hall, E. R. Collecting and Preparing Study Specimens of Vertebrates. Univ. Kansas Mus. Nat. Hist. Misc. Publ. 30:1-46.

Jennings, R. D., and N. J. Scott, Jr. 1993. Ecologically correlated morphological variation in tadpoles of the leopard frog, Rana chiricahuensis. Journal of Herpetology 27(3):285-293.

Quay, W. B. 1974. Bird and mammal specimens in fluid-objectives and methods. Curator 17(2):91-104.

Simmons, J. E. 1991. Conservation problems of fluid-preserved collections. Pp 69-89 In Cato, P. S., and C. Jones (eds.). Natural History Museums: Directions for Growth. Texas Tech University Press, Lubbock, Texas.

Simmons, J. E. 1994. Blythe Spirits: Problems and Potentials of Fluid Preservation. Paper presented at the 1992 ASIH Workshop on Collection Care and Management Issues in Herpetology and Ichthyology. gopher://kaw.keil.ukans.edu:70/00/curation/ichs_herps/1992_3.

Simmons, J. E. 1995. Storage in fluid preservatives. Pp 161-186 In Rose, C. L., C. A. Hawks, and H. H. Genoways (eds.). Storage of Natural History Collections: A Preventive Conservation Approach. Society for the Preservation of Natural History Collections, Iowa City, Iowa.

Stoddard, R. W. 1989. Fixatives and preservatives: their effects on tissue. Pp 1-25 In Horie, C. V. (ed.). Conservation of Natural History Specimens: Spirit Collections. Manchester Museum and Department of Environmental Biology, The University of Manchester.

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