Some Biological and Non-Biological Things to Consider when Sampling Amphibians in a Large-Scale Inventory and Monitoring Program

C. Kenneth Dodd, Jr.

This outline is meant to help natural resource managers, researchers, non-governmental biologists, and the interested public understand some of the biological and non-biological constraints to setting up a large-scale I & M program for southeastern amphibians. Many persons who read this will be familiar with basic amphibian biology, but others require a refresher course or are unfamiliar with amphibian life histories. The information is generalized and applies only to southeastern and US Caribbean amphibians (at least within the range of the FISC). There is usually an exception to everything discussed below. We have not included information on threats to amphibians (habitat loss and alteration, disease, exotics, climate change, xenobiotics, UVB, malformations, etc.) or the various reasons amphibians may be vulnerable to environmental problems (biphasic life cycle, skin permeability, metamorphosis, etc.). These are topics dealt with in many other publications.

Ten things to keep in mind when contemplating I & M
projects for southeastern and U.S. Caribbean amphibians.

1. There are many amphibians in the southeast/US Caribbean.

At present count, there are 83 species of salamanders and 58 species of frogs within our region. This does not include different subspecies, and certainly not genetic variants.

2. The systematic status of many species of southeastern amphibians is in a flux.

It is possible that there are a number of good, identifiable undescribed amphibians in the southeast, particularly in the genera Necturus and Siren. In addition, there is considerable debate among salamander taxonomists over what constitutes a species (in terms of genetic uniqueness). Particularly in the genera Plethodon and Desmognathus, many new "genetic" species have been described in recent years, especially in the southeastern mountains. Unfortunately, morphology and coloration may be only of limited assistance in identification; many individuals are impossible to distinguish in the field. There also are areas where considerable introgression/hybridization occurs. This has led to the recognition of species complexes (e.g., the slimy salamanders of the Plethodon glutinosus complex), or even of size-based guilds (in Desmognathus). As systematists examine other genera closely (Eurycea, Pseudotriton), the situation will probably become more complicated.

It is probably no better in the frog world, especially in the genera Pseudacris and Rana. For example, we know that bullfrogs and leopard frogs from Florida certainly look and behave differently from presumed con specifics farther north. Are sibling species involved?

3. Species and life stages are sometimes difficult to distinguish.

Even experienced herpetologists sometimes have trouble identifying not only eggs and larvae, but adults as well. There is a tremendous amount of color and morphological variation. The ability to distinguish species based on egg mass and tadpole morphology is exceptionally difficult and is an ability that is rapidly being lost, as such identification is rarely taught and the pool of knowledgeable naturalists is diminishing. There are very few current color guides to eggs and larvae, even on a local basis. Also see section 4 (below).

4. Amphibians have complex life cycles.

Terrestrial Salamanders (Plethodontidae)

The life cycle of terrestrial plethodontids takes place in a multidimensional space. We tend to think of them when they are on the surface, but surface activity may constitute only a small portion of their life cycle. Most terrestrial species probably do not have a very large home range on the actual surface (including under surface debris). However, they spend a considerable portion of their lives underground, and we really know very little about this aspect of their life history (e.g., the proportion of their time underground; the depths and range of lateral movements underground). In addition, even "terrestrial" species occasionally are arboreal. Salamanders at different life stages may remain nearly entirely underground (tiny juveniles perhaps; adults during egg deposition and mating) or on the surface (adult feeding and territoriality, environmental conditions permitting). It is by no means clear that space is used similarly by different life stages. Thus, detectability may change with life stage within a habitat.

For some terrestrial species, the eggs have never been seen, or nests have been located only with extreme infrequency. Some plethodontids may be long-lived (5-10 years).

Semi-Aquatic Salamanders (Ambystomatidae, Plethodontidae, Salamandridae)

Everything that applies to terrestrial salamanders applies to semi-aquatic salamanders in terms of surface and underground habitat use. Semi-aquatic salamanders, however, require water for reproduction. In mole salamanders and newts, breeding sites are usually standing water (ponds, ditches) free of fishes. In semi-aquatic plethodontids, breeding sites include seeps and streams from little trickle trails to good sized streams or rivers. For some species (e.g., Stereochilus) it may even include swamps.

The adults may migrate to breeding sites (mole salamanders and newts) synchronously in a quite orderly fashion, although temporally constrained to one or a few nights during the breeding season. Breeding adults and egg masses can be censussed, but we know little about what proportion of a population breeds annually, and from what area they are drawn. Males and females may not stay equal amounts of time during an entire breeding season, even when the breeding season is extended.

Stream-breeding species may live permanently in the streams (e.g., Desmognathus marmoratus), streamsides (many other Desmognathus), or at various distances from the stream (D. imitator, Gyrinophilus, Pseudotriton). Distances may be a few meters out to hundreds of meters. Breeding migrations are not synchronized. Little is known about the spatial distribution during terrestrial non-breeding times. For some species (e.g., Hemidactylium scutatum) virtually nothing is known about their lives away from streams/ditches and outside of the breeding season. For certain species, adults can be censussed streamside (e.g., D. quadramaculatus), whereas adults of other species can be found reasonably well in terrestrial habitats (e.g., D. imitator); some species can be found terrestrially as adults usually only by luck (e.g., Pseudotriton), and the adults of a few species are observed only during the breeding season (e.g., egg-brooding adult female Hemidactylium).

All eggs of semi-aquatic salamanders are laid in water, and the egg masses of some species (Ambystoma) can be censussed easily. All semi-aquatic species have larvae, which remain in a larval stage from a few months to as long as 2-3 years. Neoteny (broadly defined) occurs in a few species under favorable conditions (e.g., Ambystoma talpoideum). Larvae metamorphose and presumably take up adult habits, but nothing is known concerning dispersion for most species. Maturation can take one to many years, depending on species. Individuals of some species (Ambystoma, Notophthalmus, large Desmognathus) may live 10-15 years, or longer.

Aquatic salamanders (Amphiumidae, Cryptobranchidae, Proteidae, Sirenidae)

Little is known about the life history of most of these species, except for Cryptobranchus. For example, even the mode of reproduction (external versus internal fertilization) is unknown for Pseudobranchus. All of these species are entirely aquatic, although Amphiuma may nest on land adjacent to wetlands and move overland during extremely wet weather. The spatial use of habitat is largely unstudied except for hellbenders, who are known to have home ranges and to guard nesting sites. Fully aquatic species inhabit medium to large streams and rivers (Cryptobranchus, Necturus), mucks (Pseudobranchus, Amphiuma), and sloughs, swamps, and wet prairies (Amphiuma, Pseudobranchus, Siren). Hellbenders may live 25 or more years, but nothing is known about longevity of the others. The larvae are little known and, for the most part, rarely seen.

Frogs

All of our native southeastern frogs have a "typical" amphibian life cycle. Adults move to a breeding site, deposit eggs that hatch to larvae (tadpoles), metamorphose to juveniles, disperse, and grow until they are ready to repeat the cycle. However, the specifics of much of the life cycle (what percentage is breeding in any one year, where do juveniles go, how far do adults disperse) is still unknown for most species. Larval periods may be extremely brief (days in spadefoots) to extremely long (years in some ranids). Breeding may be synchronous (spadefoots, many ranids) or extended (e.g., bullfrogs). Even when synchronous and explosive (e.g. Rana sylvatica), the actual breeding date may extend over a period of months (say December to March) as adults wait for the right combination of environmental conditions.

It is known that adult (and juvenile?) frogs spend most of their lives away from the breeding sites, at least in most species. Individuals have been found hundreds (or even thousands) of meters from the nearest breeding sites. Frogs are often exceptionally hard to locate outside the breeding season, much less sample them. However, the terrestrial sites are extremely important to survival since individuals spend most of their lives as terrestrial predators.

In the Caribbean US, many frogs have direct development. Eggs are deposited on land in moist situations, and the "larval" period is spent within the egg. Froglets hatch as miniature adults. A few species have specialized habitats (mountain torrents, boulder caves, bromeliads) or are arboreal.

It should be obvious that while most frogs call, some species do not or they have only weak voices that do not carry far. Frog species vary the times when they call. Some species call during the day, some call at dusk and during the early evening, and some do not get going until midnight to just before dawn. Some species call only during rains, whereas others will call most evenings of the breeding season. Some frogs breed in winter (especially in the south), others breed in the spring or summer, whereas others call for an extended breeding season, even from late winter into the autumn (e.g., Hyla cinerea). Calling times and seasons also vary latitudinally.

5. In the field, detectability of amphibians is likely influenced by the following variables, to a greater or lesser extent depending on species:

• Annual cycles of reproduction
• Seasonal events (cold, drought, heat, storms) that are usually unpredictable
• Diurnal versus nocturnal activity
• Air, water and substrate temperature
• Soil moisture and rainfall
• Relative humidity
• Barometric pressure
• Cloud cover/moon brightness
• Prey availability

These variables may change daily, seasonally, or annually (e.g., El Niño versus La Niña years).

6. Species and populations occur in a landscape.

Some species are extremely localized geographically (e.g., Plethodon petraeus), whereas others are very widespread (bullfrogs). Populations may be geographically isolated to an extreme degree (e.g., Phaeognathus hubrichti; cave species; crevice-dwelling Aneides), occur very patchily in a larger landscape, occur in a metapopulation structure (e.g., toads) with considerable (or little) interchange, or occur over literally hundreds of square kilometers of marsh or deciduous forest where it is difficult to define the limits of a population (Hyla, Plethodon). Individuals may be naturally rare or exceptionally abundant. Just because something is unusual or difficult to work, do not assume it can be by-passed. Some of our most specialized species are those we know have declined or are imperiled in the southeast.

Although some populations may be huge (some hylids for example), others seem small, isolated, and vulnerable (the crevice-dwelling, cave, or ravine species). How do these species disperse? What mechanisms allow long-term persistence of small populations? Perhaps individuals move more than is recognized, or perhaps even rare immigration is sufficient to ensure genetic diversity and prevent stochastic extinction. The demography and "spatial biology" of most amphibians is still poorly understood. Even if known for a few species, the diversity of life histories suggest that generalizations about persistence will not be easily forthcoming.

An example: I monitored a striped newt population for 5 years, daily checking immigration and emigration at a small temporary breeding pond. During the latter half of the study, a great drought ensued and the population declined almost to nothing by the 5^th year. For 10 years the "pond" was dry. After 10 years (yes, the exact dates are known!), the pond finally refilled, and striped newts returned. Are they long lived? Where did the recruits come from? How did they know to come to this pond when it had been dry for so long? How would such a depression be assessed in terms of its importance to this imperiled species? These are difficult questions in trying to assess not only presence/absence, but population trends.

7. Populations may be stable or fluctuate widely.

Much of what is known concerning amphibian populations has been derived from studies of frogs and salamanders breeding in temporary ponds. The number of breeding adults and their reproductive output (larvae, metamorphs) varies to extreme proportions from one year to the next, perhaps in response to environmental and ecological conditions (weather, hydroperiod, prey availability). Some species may live in an area for years, then disappear for years only to reappear. Populations of European ranids seem to fluctuate cyclically on an 8-year cycle. On the other hand, terrestrial plethodontid populations appear rather stable from one year to the next. However, detectability may be influenced by weather (e.g., drought) even if populations are stable. Not much is known concerning the stability or fluctuation of semi-aquatic and most aquatic species and populations.

Still, we can make a wild generalization here, while keeping in mind the caveat concerning exceptions. Species that live in stable environments tend to have stable populations from one year to the next; species that live or breed in unstable or fluctuating environments tend to have populations that fluctuate to a much greater degree. Perhaps population stability can even be viewed on a gradient with environmental stability. If this is true, declines or disappearances from species living in stable environments might be more cause of concern than declines in species living or breeding in fluctuating environments.

8. Virtually nothing is known concerning emigration, immigration, and natural extinction.

It seems quite reasonable that during the course of ecological and evolutionary history, extinction and recolonization naturally occur, especially in small populations, isolated populations, or populations structured in metapopulations (sources and sinks). Yet we understand little of these processes in southeastern amphibians. The Europeans seem to have more data in attempts to understand landscape-level population changes, but their environment has been influenced by people for so long that it is difficult to separate anthropogenic from "natural" causes of extinction. In any case, remember that colonization and other forms of inter-population movements may not occur directly overland. Animals might follow sinuous topography, watersheds, streams and rivers, or even subsurface passages.

Populations of amphibians certainly experience natural turnover (i.e., recruitment, mortality), but little is known about this process or how long it takes for any North American species. Just because some individuals have the potential for considerable longevity, this does not mean that populations turn over slowly. What are the generation times for various species?

9. Amphibian sampling techniques.

There are as many ways to sample amphibians as there are amphibians. Each technique has its own underlying assumptions, biases, and limitations. Until relatively recently, these biases were unrecognized, were not discussed, or they were simply ignored. Sampling protocols have been receiving a great deal of experimental examination of late. It is unlikely that a single sampling technique can be used to sample an entire community. Some of the techniques listed below are not mutually exclusive (such as by using sweep samples within a predefined area).

Active sampling (easy to use)

• time constrained (number of observers x time sampled; catch; visual encounter)
• area constrained (using plots, transects [visual encounter surveys], habitat defined, etc.)
• sweep samples (for larvae)
• call surveys (breeding or territorial adult frogs)

Easy Passive sampling (observer need not be present; no harm to animals)

• coverboards (various sizes, shapes, configurations, materials)
• pvc pipes (in ground or on trees)
• larval litter bags
• audio data loggers (for calling frogs)

Intensive Passive sampling (labor, time, and financially expensive). These must be checked regularly, generally daily, for accurate results and to prevent mortality.

• traps (aquatic or terrestrial): funnels, bottles, minnow, wire basket, etc.
• drift fences, with pitfalls and/or funnel traps, sometimes in conjunction with pvc pipes or   coverboards.

10. The human-based constraints on sampling, inventorying, and monitoring amphibian populations on federal lands need to be considered at the outset.

These include:

• money (equipment, personnel, emergencies, meetings, data analysis, publication)
• people (PI, experienced field crews, biometricians, GIS, administrative support, field support)
• time (answers are needed, but short-term projects are ineffective. I&M needs time and patience)
• safety (two-person field crews; radios; cell phones)
• logistics (can I get there with the people and equipment in a reasonable amount of time and effort?; how many sites over what area?)
• regulations (permits; access; restrictions on research techniques; collecting)
• politics (intra-agency, federal-state, other researchers, land managers)
• bureaucracy (hiring, equipment ordering, contracts, etc.)

In our research program, we have had to deal with each of these challenges. We were prepared for some of them, and some we were not (e.g., potentially life-threatening medical emergencies involving field crews in remote locations). Each of these problems has arisen in connection with the I & M project at Great Smoky Mountains National Park. These constraints must be considered in setting up an I&M project, even if they do not have anything to do with detection probabilities or sampling per se.

In conclusion, we hope the above will give natural resource managers, scientists, and the concerned public something to think about if they contemplate an inventory and monitoring program for amphibians.

References

Adams, M. J., R. B. Bury, and S. A. Swarts. 1998. Amphibians of the Fort Lewis Military Reservation, Washington: sampling techniques and community patterns. Northwestern Naturalist 79: 12-18.

Bishop, C. A. and K. E. Pettit (eds.). 1992. Declines in Canadian amphibian populations: designing a national monitoring strategy. Canadian Wildlife Service Occasional Paper No. 76, 117 pp.

Bishop, C. A., K. E. Pettit, M. E. Gartshore, and D. A. MacLeod. 1997. Extensive monitoring of anuran populations using call counts and road transects in Ontario (1992 to 1993). in Green, D. M. (ed.), Amphibians in Decline. Canadian Studies of a Global Problem.  Herpetological Conservation No. 1:149-160.

Bonin, J. and Y. Bachand. 1997. The use of artificial covers to survey terrestrial salamanders in Québec. in Green, D. M. (ed.), Amphibians in Decline. Canadian Studies of a Global Problem. Herpetological Conservation No. 1:175-179.

Boughton, R., J. Staiger, and R. Franz. 2000. Use of PVC pipe refugia as a sampling technique for hylid treefrogs. American Midland Naturalist 144: 168-177.

Bury, R. B. and P. S. Corn. 1991. Sampling Methods for Amphibians in Streams in the Pacific Northwest. USDA Forest Service, General and Technical Report PNW-GTR-275, 29 pp.

Campbell, H.W. and S.P. Christman. 1982. Field techniques for herpetofaunal community analysis. Pages 193-200 in N. J. Scott, Jr. (ed.), Herpetological Communities, U.S. Fish and Wildlife Service, Wildlife Research Report 13, Washington, D.C.

Casazza, M. L., G. D. Wylie, and C. J. Gregory. 2000. A funnel trap modification for surface collection of aquatic amphibians and reptiles. Herpetological Review 31:91-92.

Chalmers, R. C. and S. Droege. 2002. Leaf litter bags as an index to populations of northern two-lined salamanders (Eurycea bislineata). Wildlife Society Bulletin 30: 71-74.

Corn, P. S. and R. B. Bury. 1990. Sampling Methods for Terrestrial Amphibians and Reptiles. USDA Forest Service, General and Technical Report PNW-GTR-256, 34 pp.

Corn, P. S., M. L. Jennings, and E. Muths. 1997. Survey and assessment of amphibian populations in Rocky Mountain National Park. Northwestern Naturalist 78: 34-55.

Corn, P. S., E. Muths, and W. M. Iko. 2000. A comparison in Colorado of three methods to monitor breeding amphibians. Northwestern Naturalist 81: 22-30.

Crouch, W. B., and P. W. C. Paton. 2000. Using egg-mass counts to monitor wood frog populations. Wildlife Society Bulletin 28: 895-901.

Crouch, W. B., III. and P. W. C. Paton. 2002. Assessing the use of call surveys to monitor breeding anurans in Rhode Island. Journal of Herpetology 36: 185-192.

Davis, T. 1997.  Non-distruptive monitoring of terrestrial salamanders with  artificial cover objects on southern Vancouver Island, British Columbia. in Green, D. M. (ed.), Amphibians in Decline. Canadian Studies of a Global Problem. Herpetological Conservation No 1: 161-174.

Dodd, C.K., Jr. 1990.  Line transect estimation of Red Hills salamander burrow density using a Fourier series. Copeia 1990:555-557.

Dodd, C.K., Jr. 1991. Drift fence-associated sampling bias of amphibians at a Florida sandhills temporary pond. Journal of Herpetology 25:296-301.

Fellers, G. M. and K. L. Freel. 1995. A Standardized Protocol for Surveying Aquatic Amphibians. National Park Service Technical Report NPS/WRUC/NRTR-95-01, 117 pp.

Gent, T. and S. Gibson (eds.). 1998. Herpetofauna Worker's Manual. Joint Nature Conservation Committee, Peterborough, England. 152 pp.

Gibbons, J.W. and R.D. Semlitsch. 1982. Terrestrial drift fences with pitfall traps: an effective technique for quantitative sampling of animal populations. Brimleyana 7:1-16.

Heyer, W. R., M. A. Donnelly, R. W. McDiarmid, L.-A. C. Hayek, and M. S. Foster (eds.). 1994. Measuring and Monitoring Biological Diversity. Standard Methods for Amphibians.  Smithsonian Institution Press, Washington, DC. 364 pp.

Houze, C. M., Jr. and C. R. Chandler. 2002. Evaluation of coverboards for sampling terrestrial salamanders in Georgia. Journal of Herpetology 36: 75-81.

Hyde, E. J. and T. R. Simons. 2001. Sampling plethodontid salamanders: sources of variability. Journal of Wildlife Management 65:624-632.

Jung, R. E., S. Droege, J. R. Sauer, and R.B. Landry. 2000. Evaluation of terrestrial and streamside salamander monitoring techniques at Shenandoah National Park. Environmental Monitoring & Assessment 63: 65-79.

Jung, R. E., K.E. Bonine, M.L. Rosenshield, A. de la Reza, S. Raimondo, and S. Droege. 2002. Evaluation of canoe surveys for anurans along the Rio Grande in Big Bend National Park, Texas. Journal of Herpetology 36:390-397.

Karns, D. R. 1986. Field Herpetology. Methods for the Study of Amphibians and Reptiles in Minnesota. James Ford Bell Museum of Natural History, Occasional Paper 18, 86 pp.

Lepage, M., R. Courtois, C. Daigle, and S. Matte. 1997. Surveying calling anurans in Québec using volunteers. in Green, D. M. (ed.), Amphibians in Decline. Canadian Studies of a Global Problem. Herpetological Conservation No. 1:128-140.

Lipps, K. R., J. K. Reaser, B. E. Young, and R. Ibáñez. 2001. Amphibian Monitoring in Latin America: A Protocol Manual. Society for the Study of Amphibians and Reptiles, Herpetological Circular No. 30, 115 pp.

McAlpine, D. F. 1997. A simple transect technique for estimating abundances of aquatic Ranid frogs. in Green, D. M. (ed.), Amphibians in Decline. Canadian Studies of a Global Problem. Herpetological Conservation No. 1:180-184.

Mitchell, J. C. 1998. Amphibian Decline in the Mid-Atlantic Region: Monitoring and Management of a Sensitive Resource. U.S. Department of Defense Legacy Resource Management Program, 144 pp.

Mitchell, J. C. 1998. Guide to Inventory and Monitoring of Amphibians on Fort A.P. Hill, Fort Belvoir, Marine Corps Base Quantico, and Prince William Forest Park, Virginia. U.S. Department of Defense Legacy Resource Management Program, Supplement 1, 66 pp.

Mitchell, J. C. 1998. Guide to Inventory and Monitoring of Streamside Salamanders in Shenandoah National Park, Virginia. U.S. Department of Defense Legacy Resource Management Program, Supplement 2, 29 pp.

Mitchell, J. C. 2000. Amphibian Monitoring Methods & Field Guide. Smithsonian Conservation Research Center, Front Royal, Virginia. 56 pp.

Monti, L., M. Hunter, and J. Witham. 2001. An evaluation of the artificial cover object (ACO) method for monitoring populations of the redback salamander Plethodon cinereus. Journal of Herpetology 34: 624-629.

Olson, D. H., W. P. Leonard, and R. B. Bury (eds.). 1997. Sampling Amphibians in Lentic Habitats. Northwest Fauna No. 4, 134 pp.

Pauley, T. K., and M. Little. 1998. A new technique to monitor larval and juvenile salamanders in stream habitats.  Banisteria 12:3236.

Pearman, P. B., A. M. Velasco, and A. López. 1995. Tropical amphibian monitoring: a comparison of methods for detecting intra-site variation in species' composition.  Herpetologica 51: 325-337.

Sargent, L. G. 2000. Frog and toad population monitoring in Michigan. Journal of the Iowa Academy of Science 107: 195-199.

Smit, G., A. Zuiderwijk, and A. Groenveld. 1998. A national amphibian monitoring program in the Netherlands. SEH, Le Bourget du Lac, France. Pp. 397-402.

Sorensen, K. 2003. Trapping success and population analysis of Siren lacertina and Amphiuma means. M.S. thesis, University of Florida, Gainesville.

Taylor, A., G. Watson, G. Grigg, and H. McCallum. 1996. Monitoring frog communities: an application of machine learning. AAAI/IAAI 2: 1564-1569. [8th Innovative Applications of Artificial Intelligence Conference, Portland, Oregon]

Vogt, R. C. and R. L. Hine. 1982. Evaluation of techniques for assessment of amphibian and reptile populations in Wisconsin. Pages 201-217 in N. J. Scott, Jr. (ed.), Herpetological Communities, U.S. Fish and Wildlife Service, Wildlife Research Report 13, Washington, D.C.

Waldron, J. L., C. K. Dodd, Jr., and J. D. Corser. 2003. Leaf litterbags: factors affecting capture of stream-dwelling salamanders. Applied Herpetology, in press.

Wilson, C R. and P. B. Pearman. 2000. Sampling characteristics of aquatic funnel traps for monitoring populations of adult rough-skinned newts (Taricha granulosa) in lentic habitats. Northwestern Naturalist 81: 31-34.

Wilson, J. J. and T. J. Maret. 2002. A comparison of two methods for estimating the abundance of amphibians in aquatic habitats. Herpetological Review 33:108-110.