Appendix 4 Preparing Amphibians as Scientific Specimens Roy W. McDiarmid Introduction Documentation of the species recorded in biodiversi- ty and inventory projects frequently involves prepa- ration of voucher specimens, which are extremely important for the credibility of such studies (see Chapter 5). In addition, these specimens and their ac- companying field notes are valuable sources of scien- tific information used in systematic, biogeographic, evolutionary, and ecological studies. Most vouchers are preserved specimens from the study site, although photographs of individuals or tape-recorded calls of frogs are sometimes substi- tuted. To be useful as vouchers and to facilitate iden- tifications of species, preserved specimens must be well prepared and well documented. Poorly prepared or improperly documented specimens are of little sci- entific value and usually not worth the effort of prep- aration. In this section I review the steps involved in the proper preparation of amphibians as scientific study specimens (for additional discussion, see Pisani 1973; Kams 1986; Simmons 1987). Documentation Field notes should be written on 100% cotton rag paper with permanent ink. Most field biologists re- cord field notes on loose-leaf paper stored in a three- ring binder. Although some people argue that loose-leaf binders are inferior to bound volumes be- cause pages may be lost, this is not a problem if a person is careful. A loose-leaf format provides flexi- bility and allows for easy insertion of extra pages, maps, lists, and other important documents. Because bound books have a fixed size, they may contain too many pages for short trips and too few pages for 2^ 290 APPENDIX 4 ?/ ^Jaii, r^ /fH i/e-?c 2.UC a. : Mrri^ffO fec/'^^t^-C K'Mxz.o-p.t^s / M-?/ina. '??f^ . OH ^'p ?a-ricK.- /^?itr: p's-?'tj , ?C '/? ?f/ dasiroa. f7C8f /^?gnr? e nei-h Jrs. S^. r70?L ^^? /) ?J^ T^? t-i?^^f ?. Tjv^ ^Ara^>/acuc?is/5 Uko/-kxJtt^ Figure 30. Sample page from a field notebook. Preparing Scientific Specimens 291 long ones. Nevertheless, I recommend against taking a bound volume with original notes into the field on a subsequent trip because of the possibility of loss. This is not a problem with a loose-leaf notebook, be- cause original pages can be left behind. Loose-leaf pages can easily be bound into annual or trip-related volumes for permanent archives. E*reserved specimens should be accompanied by the minimum data discussed in Chapter 5: locality, date and time of collection, collector, sampling method, habitat, and other locality-related informa- tion. Other specimen-related information?such as color notes and references to photographs, tape re- cordings, and tissue samples?should be recorded in the field notes (Fig. 30). Each reference to a voucher specimen in the notes should include the field num- ber and temporary field identification (e.g., frog; Rana sp,; small brown salamander). The number used in the field notes is written on a field tag and attached to the specimen (see below). Most field biologists use field numbers that include their mitials followed by a number (e.g., RWM 19503), but any distinct numbering scheme will suf- fice (simpler is better). Field tags should be rela- tively small and of sturdy paper stock (Kams [1986] suggested Resistall Index Bristol, 100% rag, 110 lb. wt. paper, available Ironi University Products; see Appendix 6), Field numbers should be written on the tag in f)ermanent ink. Printed tags with consecutive numbers and other identifiers are available commer- cially. Ahhough printed tags are relatively expensive (U.S. $20-S100 per 1,000 tags. National Tag Com- pany), I recommend them because they save valu- able field time and prevent double numbering. Tags of cloth or plastic can be used if paper tags are not RWM 19503I Figure 31. Numbered field tag and attached thread. Letters are the initials of the collector, and the numbers are consec- utively printed ones. Note the location of the loop hitch, which holds the tag, and the knot, which holds the tag away from the animal. available, but metal tags should be avoided because they sometimes corrode, and sharp edges can dam- age fragile specimens. Field tags should be strung with a fme white linen thread as shown in Figure 31. Colored thread should not be used because the dye may dissolve in preservative and discolor the specimens. Specimen processing Immature and adult amphibians collected as vouch- ers should be maintained in plastic bags until pro- cessed. Eggs and larvae require immediate attention, and I discuss them separately. When I capture a spec- imen, I usually place a numbered plastic tag in the bag with it and use that number to reference associ- ated data in a small pocket notebook. I do not write data on paper tags and place them in die bag with the specimen because these quickly become wet and illegible. As soon as possible, I transfer pertinent data for each specimen from the pocket notebook to permanent field notes and assign a field number to the specimen. The plastic field tags can be washed and reused. I cry to process all specimens as quickly as possible to avoid loss of individuals and confusion of associated data. Dead amphibians generally do not preserve well and make poor voucher specimens. In some situations specimens may be frozen for later processing. However, Scott and Aquino-Shuster (1989) showed that amphibians frozen before preser- vation are softer and grayer than specimens that were not frozen, often have twisted toes, and lose their epidermis. Because some of the traits affected are important for species identification (e.g., color- ation, toe and webbing characters, skin texture), I strongly recommend that specimens not be frozen. Procedures for Killing Amphibians should be killed as quickly and hu- manely as possible in a way diat leaves them in a re- laxed condition. The most efficient method for killing adult amphibians is to immerse them in a so- lution of Chloretone. A Chloretone solution is made by dissolving a small amount (1 teaspoon) of hy- drous chlorobutanol crystals in a Uter of water. Note that chlorobutanol crystals (and hypodermic syrin- ges) are controlled substances in some countries, and investigators may need permits to obtain t?iem. Be- 292 APPENDIX 4 cause chlorobutanol dissolves slowly, I often carry a small container of stock solution (95% ethyl alcohol saturated with chlorobutanol) and prepare a Chloretone solution in the field by adding a few mil- liliters of stock solution to 750 m\ of water in a wide- mouthed liter bottle. I prepare a Chloretone solution as soon as I arrive at a field site so that it is ready when needed. Such a solution is effective for about 1 to 2 weeks (with heavy use) and then gradually loses its strength. Species have noticeably different responses to Chloretone solutions. Some amphibians die rather quickly (5 min); others may take longer (10-15 min). The solution often increases in effectiveness, probably from the addition of various skin secretions from am- phibians killed in the solution. If amphibians do not die after 20 to 30 minutes in the solution, additional chloro- butanol should be added or, preferably, a new solution prepared. Amphibians should not be left in the solution very long after they die, as they become rigid and diffi- cult to position for fixation. If Chloretone is not available, amphibians may be killed by drowning in warm water (43''-47?C) or in weak alcohol (15%-25%) solutions (Pisani 1973). Pith- ing may be effective, but it often damages the speci- men, especially the skuU. Benzocaine-containing gels, sold in most pharmacies as toothache medication, can be used as an alternative to Chloretone (Altig 1980). A small amount smeared on the head of an amphibian kills it wi?iin a few minutes, and the specimen is com- pletely relaxed. Freezing is not recoinmended as a method for killing (Scott and Aquino-Shuster 1989). Preservatives Preserving amphibians for scientific study is usually a two-step process. Initially, specimens are fixed in an appropriate preservative, and then they are trans- ferred to an alcohol solution for storage. The most common general fixative for amphibians is formalde- hyde. At room temperatures, formaldehyde is a gas; under pressure, it polymerizes to a solid called paraformaldehyde. An aqueous solution of formalde- hyde is easier to handle than the gas and has been used as a histological fixative for about 100 years (Fox and Benton 1987). Formaldehyde is commer- cially available as a liquid that is made by dissolving formaldehyde gas in water as a 37% to 40% solu- tion. A formaldehyde solution of this strength is equivalent to 100% formalin (formaldehyde diluted with water). One part of this full strength formalin is diluted with nine parts water to make a 10% solution of formalin. A 10% solution is the standard for specimen fixa- tion in the field. A fixed specimen retains whatever position it had when placed in the formalin. This sta- bilization (fixation) is effected by cross-linkage of the formalin with protein end groups in the tissue. This process stops autolysis and prevents further tis- sue deterioration. Although the rate of fixation varies with the size of the specimen and probably with the nature of the tissues, most small amphibians set in a few hours; larger specimens require much longer. Fixation generally is considered to be irreversible, but Pearse (cited in Taylor 1977) found that many of the cross-linking bonds are reversible with the sim- ple process of washing; others are not. Because washing under certain situations may reverse fixa- tion, I recommend that amphibians not be soaked in water prior to transfer to ethyl alcohol. Because formaldehyde is a gas, a formalin solu- tion exposed to air decreases in strength and be- comes more acidic. It is especially important, therefore, to use freshly prepared 10% solutions to fix specimens. I recommend replacing formalin in trays after preserving 50 to 100 specimens of various sizes. Old formalin can be used in the field as a stor- age solution for specimens after fixation, or it should be properly discarded in keeping with regulations of the country where the investigator is working. Formaldehyde oxidizes to form formic acid, which, after a time, will decalcify bone and discolor specimens. Specimens fixed in formalin that is alka- line, in contrast, tend to become transparent, or "cleared" (Taylor 1977). Because d?calcification, ex- cessive discoloration, and clearing are undesirable, most field herpetologists buffer formalin to maintain the pH as close to neutral as possible. Fox and Ben- ton (1987) indicated that an optimum pH for forma- hn fixation is 7.2. Although borax has been suggested as a suitable buffer (Pisani 1973), Taylor (1977) argued against its use because specimens tend to clear in borax-buffered formalin; instead he recommended calcium carbonate. In the field I have used powdered magnesium carbonate with good re- sults (about 1/2 teaspoon per liter of 10% formalin). To buffer formalin used in the museum, I add 4.0 g of sodium phosphate monobasic and 6.5 g of sodium phosphate dibasic (anhydrous) to each liter of 10% formalin. In the museum, I prepare formalin with distilled water, but in the field I use whatever water is avail- able. Field researchers should be aware, however. Preparing Scientific Specimens 293 that stream or pond water may be quite naturally acidic or basic, and, therefore, formalin made with such water may require additional treatment. The use of rainwater often alleviates the pH problem, but even rainwater can be acidic. If the pH of water at a field site is potentially a problem, then it may be de- sirable to check the pH of the solution with indicator paper. I use a pH meter hi the museum. Occasionally in remote areas, one purchases bad formalin (i.e., it does not fix animals as well as expected). I suspect that such formalin has been oxi- dized extensively and probably contains consider- able formic acid. Workers need to be aware of this problem and to test the pH of all formalin solutions. I repeatedly examine fixed materials in the field to avoid problems of poor fixation. Formaldehyde fixa- tion is a complicated process and can be influenced by many extraneous factors. If an investigator is ob- taining the desired results, I recommend that he or she continue the process. In the absence of desired results, I recommend starting over with firesh solu- tions. If the problem persists, help from a chemist or a museum curator should be sought. As most heipetologists know, formaldehyde is bri- tating to the eyes, upper respiratory passages, and skin. Some people develop an allergic reaction to for- malin and must wear rubber gloves to prevent skin rashes. Formaldehyde also has been reported to be a potential carcinogen (Simmons 1987); thus, it should be used only in well-ventilated areas. Fortunately, most field situations fall into this category. If formaldehyde is not available as a fixative, 70% ethyl alcohol can be used. I do not recommend other kinds of alcohol (e.g., methyl, isopropyl, rubbing). Fixatives such as formalin-acetic-alcohol (FAA), glu- taraldehyde (Taylor 1977), and Bouin's solution (es- pecially good for histol?gica! work) are more difficult to prepare but also can be used. Formalin so- lutions also can be prepared from parafonnaldehyde (see recommendations in Huheey 1963; Pisani 1973; Saul 1981). Fixation Once an amphibian is dead and relaxed, it can be fixed in a preserving tray in 10% formalin. Typi- cally, I use a plastic refrigerator tray (33 x 21 x 6 cm) with a tightly fitted lid. I line the bottom with a white paper towel soaked with 10% formalin (dye from colored towels will discolor specimens). Each specimen is positioned in the tray in a way that wiU facilitate measurement and examination of key fea- tures on preserved specimens and that also will allow for more-effective storage and hence long- term maintenance of the specimens. I position fi-ogs so that their limbs are drawn in next to the body and flexed in a natural way; fingers and toes are straight- ened and spread to display webbing (Fig. 32). I do not recommend the positioning illustrated by Pisani (1973: plate 1) or Kams (1986: fig. 24), in which one or both hind limbs of a frog are extended posteri- orly. In my experience extended limbs easily become tangled with other specimens and tags as ani- mals are removed from a storage jar. Often the q)i- derniis is scraped excessively and the hind feet damaged. From a practical perspective, frogs with ex- tended limbs require more storage space (fewer frogs per container). They also require closer monitoring; the toes and webbmg on the hind feet are particularly sub- ject to desiccation and may be the first structures ex- posed to air as the alcohol level in the jar drops. Most salamanders are laid out straight with the limbs pointing forward parallel to the body. The hands and feet are arranged with palmar surfaces down and toes spread, and the tail is straight (Fig. 32). Caecilians are preserved with the body straight or in a flat loop of a size appropriate for the containers used in the collection (Fig. 32). I preserve caecilians with their mouths held open by a small stick or piece of paper towel; this practice facihtates later examination of tooth arrangement. Formalin penetrates the body cavity of small am- phibians rather quickly, so injection is not necessary. Large frogs (adults of many species of Bufo, Rana, and Leptodactylus), salamanders (sirenids, crypto- branchids, and adults of larger species of Ambystoma), and caecilians may require injections of formalin into the body cavity and larger muscle masses. Care should be taken not to distend the body by injecting too much formalin and not to introduce air into the body cavity. Frogs distended with air float in the hardening solution, so some areas of the body are not covered with fixadve. It also is import- ant to keep track of individuals in the tray so that the correct field tag can be associated with the appropri- ate individual after it has hardened. I sometimes lay the tag across the specimen in the tray. Once the floor of the tray is covered with specimens, I blanket them with a second paper towel wet with formalin and carefully fill the tray with formalin to about one third its depth. After a few hours, most specimens will have har- dened enough to maintain their shape. This is the 294 APPENDIX 4 Figure 32. Specimens of frogs, salamanders, and a caeciiian preserved in the recommended postures (see text for explanation). time to attach the field tag. Soine herpetologists prefer to attach tags to specimens prior to position- ing them in the hardening tray to ensure that tags and specimens do not get mixed. I have found, however, that specimens are more difficult to posi- tion for hardening with the tag attached. The tag is tied with a square knot above the knee on the right rear leg of frogs and large salamanders, and around the neck of small salamanders and caeci- lians; ends of the thread are trimmed. The speci- mens are transferred to a hardening jar, where they remain immersed in 10% formalin for at least 4 days or, preferably, for the remainder of the field season. Eggs and Larvae Amphibian eggs and larvae usually require special treatment. They are easily damaged, especially dur- ing collecting, if not handled carefully. As a resuU, when collecting eggs or tadpoles I usually preserve some in the field and take others back to camp alive. I place single, short strings or small clumps of aquatic eggs directly into small bottles or vials of freshly prepared 10% buffered formalin that I carry with me in the field for that purpose. I usually place terrestrial eggs, eggs adhering to leaves, and larger egg masses (e.g., those of some species ofAmby- stoma), into plastic bags and carefully transport them back to the work area for examination, rearing (see below), and preservation in suitably sized con- tainers. I put some larvae directly into formalin as they are collected, and carry others back to camp in plastic bags. Because eggs and larvae, especially of anurans, contain much more water than adults con- tain, they seemingly require a larger volume or slightly stronger concenfration of fixative initially than do adults. I sometimes carry a small amount of pure formalin into the field to meet such needs. On reluming to camp or the lab, I routinely sort and transfer all preserved eggs and larvae to larger containers of fresh 10% formalin. During the sorting process, I identify larval morphotypes, write color notes, and take photographs. At this time I also de- cide whether to maintain some of the live eggs until Preparing Scientific Specimens 295 hatching or to rear some of the live tadpoles through metamorphosis. 1 often preserve a few eggs from a clutch and place the reinainder in an appropriate container to continue their development. With lar- vae, I often return to the site and collect additional specimens for rearing (see below). After sorting the specimens, I write field notes and assign tags to the larval samples. Tadpoles that have been in an adequate volume of fresh formalin for 24 hours arc well fixed and can be transferred with their field tags to smaller vials for storage and eventual transport. A tag should never be tied to a tadpole or to a small salamander or caecilian larva; tags may be attached to larger salamander and caeci- lian larvae if tagging does not damage the speci- men. I prefer to sort samples into morphotypes in the field and tag them accordingly. However, if storage space or tags are limited, al! specimens collected at a single site on a single day can be maintained as a single sample. Samples collected at different localities or sites or on different days must not be mixed. Because the eggs and larvae of many species are unknown, efforts should be made to associate each with its respective adult form. If time per- mits, an investigator can obtain and rear eggs from known adults or can rear unknown larvae through metamorphosis in the field. Aliquot samples representative of the eggs and larvae from a single known sample can be preserved at appropriate intervals during development. Such developmental series are very useful in under- standing the ecology of species and may contrib- ute significantly to investigations of the evolution of morphological traits in amphibians. Containers suitable for rearing tadpoles can be set up in camp, and tadpoles of specific morphotypes can be reared. I usually maintain each sample (species) separately in water taken from its habitat and, if possible, feed the tadpoles natural-occurring food. I use artificial food (e.g., tropical fish food, trout chow, rabbit pellets) if nat- ural foods are not easily obtainable. Periodic sampling of tadpoles from the same aquatic habi- tat sometimes is more efficient than rearing them. Tadpoles sampled from populations maintained ar- tificially in the field or laboratory should be given separate numbers and cross-referenced in field notes to the field number of the original sample. Date, time of sampling, and rearing conditions (e.g., container size, temperature, food) for each sample should be recorded. Special preparations Sometimes it is desirable to prepare specimens for osteol?gica! study. Because of distortion from dry- ing, cleared and stained preparations (see below) are preferable for most species of amphibians. However, dry preparations of larger species are suitable and sometimes preferred. Simmons (1986) described a method for making osteological preparations from al- coholic specimens. I recommend his approach, al- though it is time-consuming. If a specimen is prepared as a dry skeleton in the field, the selected individual should be weighed and measured before it is put into Chloretone. Skinning should follow the steps outlined by Simmons (1986), except that no parts of the skeleton should be disassociated. The sex of the specimen should be determined when the animal is eviscerated, and stomach contents should be recorded. If the latter seem interesting, they should be preserved and referenced with the same field number as the specimen. To make an osteological specimen, the preparator removes most of the muscle mass, being careful not to cut the pectoral girdle or hyoid apparatus, and at- taches a field tag to the leg once it is mostly free of tissue. The specimen is then wrapped into a compact but loose ball with string and hung in a dry place, preferably within a screened enclosure or cage to dis- courage flies and animals that may carry off a car- cass. If the specimen does not dry quickly, it can be immersed in alcohol for a few hours and dried again. It should not be put in formalin, because dermestid beetles, which are used to clean skeletal material, will not eat formalin-treated carcasses. Putting the carcass in a small cheesecloth bag will help to ex- clude flies and to maintain all skeletal elements in a single package. Once a carcass is thoroughly dry, it can be packed in its cheesecloth bag for shipment back to the museum. Simmons (1987) provided many references to techniques for preparing skele- tons and cleared and stained specimens. Packing and shipping At the end of a field season or work at a site, all pre- pared specimens should be checked to see that field tags are attached and that no problems with associ- ated data remain. If live material is to be taken to the next site or back to the museum for continued rear- ing (a job that is sometimes difficuh but potentially 296 APPENDIX 4 worth the effort), then a sample should be preserved at the last possible moment to ensure that representa- tive material at that stage of development is avail- able for study. Preserved specimens are sorted by shape, size, and method of preparation. Dry skeletal preparations should be packed in dry containers (small cans or du- rable cardboard boxes) with lots of padding. Vials containing eggs and larvae are grouped by size and wrapped with packing material or strips of paper towel. Vials of similar size are packed tightly in plas- tic jars, cans, or other appropriate containers with paper or cardboard dividers, as necessary. I pack vials in scalable contamers (cans or plastic jars) rather than in cardboard boxes because of the possi- bility of leaks or breakage during transport. If card- board boxes are used, they should be sealed in plastic bags. Wet specimens should be loosely wrapped in paper towels or cheesecloth for protec- tion and easier packing. Many small packages, each containing a few specimens, seem to be better than fewer, larger packages. If the collecting site is near the laboratory, so that collections can be returned from the field by car, then transporting wrapped specimens in trays with formalin works well if the trays are kept flat. Small specimens can be left in for- mahn in sealed containers. If collections have to be shipped from the field site to the laboratory or taken as baggage on an air- plane, then different packing is required. Weight and leakage are primary considerations, and plastic con- tainers are much better than glass ones. Wet speci- mens should be wrapped in cheesecloth or paper towels and placed in a plastic bag. When the bag is about one third full of one or more equal-sized bun- dles, just enough formalin to soak the packages should be added, and the bag tied securely or closed with a rubber band. After the bag is checked for leaks, it is placed in a second bag that also is tied. Bags of comparable size can be packed snugly in plastic jars, preserving trays, or other sealable con- tainers, which are then taped shut. These containers can be packed into shipping boxes or fiberboard drums designed for shipping wet materials. An ad- dress label should be placed on the inside as well as on the outside of each shipping container. Some- times I hand-carry small containers of fragile speci- mens or vials of eggs and larvae. I have carried half-liter plastic jars filled with small frogs and sala- manders packed between layers of cheesecloth wet with formalin, with good success. The formalin in these containers can be replaced with 70% ethyl alco- hol for transport on an airplane. If I know that a trip is going to be short (< 12 hr), and I am hand-carry- ing specimens, I sometimes replace the formalin in vials with water. This ensures that formalin will not be spilled during transport. I refill the vials with fresh 10% formalin immediately on reaching my des- tination. Larvae need to be shipped in vials or sim- ilar containers with some liquid and should never be wrapped in cheesecloth. With careful packing and common sense, important materials will arrive un- harmed. Field equipment and supplies One or more good-quality headlamps, along with batteries and replacement bulbs, are essential for nighttime survey work. I rely on a plastic headlamp with four D-size batteries. I prefer D-cells over a sin- gle 6-volt battery because D-cells generally aie more readily available and easier to pack. Some people use rechargeable D-cells, but in my experience, they do not last very long and do not provide the service that an alkaline cell gives. Another option is the more powerfiil, rechargeable miner's lamp with a wet-cell (motorcycle type) battery. A miner's lamp gives a brighter beam and, though initially more ex- pensive, may be less expensive in the end, because the batteries can be recharged hundreds of times. However, like nickel-cadmium dry cells, the batter- ies require electricity or solar panels for recharging; also they are hard to pack and burdensome to main- tain during long periods of disuse. The following Ust of equipment and supplies is provided as an aid to preparing for fieldwork. Sources for these supplies can be found in Appen- dix 6. Collecting Equipment. Headlamp and batteries; re- placement bulbs; small flashlight and batteries; as- sorted plastic bags; cloth bags; temporary plastic field tags; thermometers; compass; altimeter; ma- chete and file; pocket knife; insect repellent. Equipment for Observing, Studying, and Measur- ing. Measuring tape; marking flag; calipers; plas- tic rulers (10- and 30-cm); spring scales (10-, 100-, and 500-g); small scissors; hand lens; cam- era and film; tape recorder, microphone, batteries, and tape; binoculars. Equipment for Recording Data. Notebook and waterproof paper; small pocket notebook; pens Preparing Scientific Specimens 297 and ink; pencils; Sharpie indelible-ink marking pen; field tags; waterproof paper for duplicate tags; thread; scissors (for paper); cigarette lighter; candles for light when writing. Equipment for Larval Sampling. Dipnets (large and small); spare net bags; small-mesh strainers; assorted vials and small jars; large-gauge pipette; rearing containers (plastic food containers with lids); plastic bags; artificial food (trout chow, tropi- cal fish flakes); plastic spoon. Preserving Equipment. Forceps (long and needle- nosed); dissecting scissors; scalpel and blades; single-edged razor blades; needle and thread; pre- serving trays; wide-mouth jar for Chloretone; sy- ringes (1-, 10-, and 30-cc); syringe needles (vari- ous sizes); hardening and storage jars; paper towels. Chemicals. Formalin (full strength); buffer (mag- nesium carbonate); chlorobutanol or Chloretone (saturated solution); alcohol (full-strength etha- nol). Containers. Assorted plastic jars (0.5- to 3.2- liter); shipping containers; bucket or Liquipak (a watertight fiber drum for shipping wet materi- als). Pacldng Supplies. Fiber tape; string; rubber bands; cheesecloth; heavy plastic bags; scissors (for cheesecloth); paper towels; mailing labels; Sharpie indelible-ink pen.