Monitoring Amphibians in 

Great Smoky Mountains 

National Park

U.S. Department of the Interior

U.S. Geological Survey

Circular 1258

By C. Kenneth Dodd, Jr.
Photographs by: Author unless otherwise noted, between 1998 and 2002. All animals were photographed 

within Great Smoky Mountains National Park.

Color illustrations by: Jacqualine Grant, Cornell University.

Layout: Patsy Mixson

Graphic Design: Jim Tomberlin
Monitoring Amphibians in Great Smoky 

Mountains National Park

By C. Kenneth Dodd, Jr.

U.S. Geological Survey Circular 1258

U.S. Department of the Interior

U.S. Geological Survey
The use of firm, product, trade, and brand names in this report is for identification 

purposes only and does not constitute endorsement by the U.S. Geological Survey.

Tallahassee, Florida

2003

For additional information write to:

C. Kenneth Dodd, Jr.

Florida Integrated Science Center

U.S. Geological Survey

7920 N.W. 71st Street

Gainesville, Florida 32653

For additional copies please contact:

U.S. Geological Survey

Branch of Information Services

Box 25286

Denver, CO 80225-0286

Telephone: 1-888-ASK-USGS

World Wide Web: http://www.usgs.gov

Library of Congress Cataloging-in-Publication Data

Dodd, C. Kenneth.

Monitoring Amphibians in Great Smoky Mountains National Park / by C. Kenneth Dodd, Jr.

p. cm. — (U.S. Geological Survey circular ; 1258)

Includes bibliographical references.

ISBN 0-607-93448-4 (alk. paper)

1. Amphibians — Monitoring — Great Smoky Mountains National Park (N.C. and Tenn.) 2. Great Smoky 

Mountains National Park (N.C. and Tenn.) I. Title. II. Series.

QL653.N8D64 2003

597.8’09768’89—dc22

2003061257

U.S. DEPARTMENT OF THE INTERIOR

GALE A. NORTON, Secretary

U.S. GEOLOGICAL SURVEY

CHARLES G. GROAT, Director
III

Contents

1 Abstract 43 Choosing Sampling Sites

2 Introduction 44 Sampling Watersheds

4 Acknowledgments 44 Sampling Streams

5 How to Use this Guide 44 Sampling Locations

6 Amphibians of the Great 

Smoky Mountains

45

45

Sampling Techniques and Protocols

Active Sampling

6 Species Richness 53 Easy Passive Sampling

7 Habitats and Distribution 60 Intensive Passive Sampling

11 Salamanders 63 Data Handling 

20 Frogs 63 Field Data

20 Ponds 64 Spreadsheets and Databases

23 Woodland Pools 64 Analysis and Software 

23 Grassy Ditches, Pools, and Rivulets 66 Equipment and Training 

23 Streams and Rivers 67 Biosecurity and Disease

26 Other Breeding Sites 67 Biosecurity Protocol

26 Life History 68 Disease Protocols 

29 Areas of Particular Amphibian 

Species Richness

69

69

Methods

Live and Sick Amphibians

29 Identification 69 Dead Amphibians 

29 Salamanders 70 Labels

30 Frogs 70 Mailing

33 Additional Information 71 Quarantine of Amphibians 

34 Monitoring Programs 71 Malformations 

34 Why Monitor Amphibians? 72 Conclusions 

35 Things to Consider During Planning 74 Summary

40 Species and Locations to Monitor 76 References on Inventorying and 

Monitoring Amphibians
IV

Appendixes

Plates

82 Appendix I. Location of selected wetland sampling sites in Great Smoky Mountains 

National Park

83 Appendix II. Example of a field data sheet prepared for amphibian surveys in 

Great Smoky Mountains National Park

84 Appendix III. Sampling equipment

87 Appendix IV. Guidelines for building and operating remote field recorders 

(automated frog call data loggers)

Amphibians

100 1A. Marbled Salamander (Ambystoma opacum). Stupka=s Sinkhole Pond, Cades 

Cove.

100 1B. Spring Salamander (Gyrinophilus porphyriticus). Andrews Bald.

101 2A. Shovel-nosed Salamander (Desmognathus marmoratus). Mill Creek, Cades 

Cove.

101 2B. Jordan=s Salamander (Plethodon jordani). Clingmans Dome.

102 3A. American Toad (Bufo americanus). Cades Cove.

102 3B. Wood Frog (Rana sylvatica). Gregorys Cave, Cades Cove.

Habitats

104 4A. View southwest from Clingmans Dome.

104 4B. Mt. LeConte.

105 5A. Top of Andrews Bald, view toward Thunderhead Mountain.

105 5B. Clingmans Dome (highest point in the Great Smokies).

106 6A. Cades Cove, view to southeast.

106 6B. Cades Cove, view toward Gregorys Bald (hidden in clouds).

Larvae and Tadpole Mouthparts

108 7A. Spotted Salamander (Ambystoma maculatum).

108 7B. Marbled Salamander (Ambystoma opacum).

108 7C. Mole Salamander (Ambystoma talpoideum).

109 8A. Hellbender (Cryptobranchus alleganiensis).

109 8B. Spotted Dusky Salamander (Desmognathus conanti).

109 8C. Imitator Salamander (Desmognathus imitator).

109 8D. Shovel-nosed Salamander (Desmognathus marmoratus).

109 8E. Seal Salamander (Desmognathus monticola).

110 9A. Ocoee Salamander (Desmognathus ocoee).

110 9B. Black-bellied Salamander (Desmognathus quadramaculatus).

110 9C. Santeetlah Salamander (Desmognathus santeetlah).

110 9D. Three-lined Salamander (Eurycea guttolineata).

110 9E. Junaluska Salamander (Eurycea junaluska).

111 10A. Long-tailed Salamander (Eurycea longicauda).
V

111 10B. Cave Salamander (Eurycea lucifuga).

111 10C. Blue Ridge Two-lined Salamander (Eurycea wilderae).

111 10D. Spring Salamander (Gyrinophilus porphyriticus).

111 10E. Four-toed Salamander (Hemidactylium scutatum).

112 11A. Common Mudpuppy (Necturus maculosus).

112 11B. Eastern Red-spotted Newt (Notophthalmus viridescens).

112 11C. Mud Salamander (Pseudotriton montanus).

112 11D. Black-chinned Red Salamander (Pseudotriton ruber).

113 12A. Northern Cricket Frog (Acris crepitans).

113 12B. American Toad (Bufo americanus).

113 12C. Fowler=s Toad (Bufo fowleri).

114 13A. Eastern Narrow-mouthed Toad (Gastrophryne carolinensis).

114 13B. Cope=s Gray Treefrog (Hyla chrysoscelis).

114 13C. Spring Peeper (Pseudacris crucifer).

115 14A. Upland Chorus Frog (Pseudacris feriarum).

115 14B. American Bullfrog (Rana catesbeiana).

116 15A. Northern Green Frog (Rana clamitans).

116 15B. Pickerel Frog (Rana palustris).

117 16A. Northern Leopard Frog (Rana pipiens).

117 16B. Wood Frog (Rana sylvatica).

117 16C. Eastern Spadefoot (Scaphiopus holbrooki).

Figures

3 1. Map of Great Smoky Mountains National Park, North Carolina and 

Tennessee

4 2. Map showing location of U.S. Geological Survey sampling sites in Great Smoky 

Mountains National Park, 1998 to 2001

7

3-26. Photos of:

3. Spruce-fir forest at Indian Gap

8 4. Deciduous forest at Lynn Hollow

9 5. Hemlock forest at Chinquapin Knob

10 6. Coarse woody debris in Cove forest at Roaring Fork

11 7. Oak-pine forest

14 8. Middle Prong at Tremont

15 9. Ideal habitat for Hellbenders in Lower Abrams Creek

15 10. Small stream in unnamed tributary to Falls Branch

16 11. Medium-sized stream in normal flow at Roaring Fork

16 12. Medium-sized stream in high water at Whiteoak Flats Branch

17 13. Entrance to Gregorys Cave

18 14. Rimstone pools and cave pool at Gregorys Cave

18 15. Rock face near Double Gap

19 16. Former trout pond mucky habitat in Cataloochee

19 17. Seep at Big Spring Cove
VI

20 18. Gum Swamp at Cades Cove in high water

20 19. Gum Swamp at Cades Cove when dry

21 20. Gourley Pond at Cades Cove in high water

21 21. Gourley Pond at Cades Cove when dry

22 22. Methodist Church Pond at Cades Cove

22 23. Beaver pond at Big Cove

23 24. Woodland drainage pool at Cane Creek

26 25. Grassy pool at Cades Cove

27 26. Chilhowee Lake at mouth of Abrams Creek

30 27. Illustration of body morphology of a salamander larva: pond form; stream form

31 28. Illustration of body morphology of a tadpole

32 29. Illustration of oral disc (mouthparts) of a tadpole

41

30-34. Photos of:

30. Biologist sampling with sweep net in Stupkas Sinkhole Pond at Cades Cove

42 31. Biologist looking for tadpoles at Shields Pond in Cades Cove

45 32. Turning logs in time-constrained survey at Beech Flats

46 33. Terrestrial time-constrained survey in thickly vegetated habitat at Balsam 

Mountain

46 34. Dip netting for salamander larvae in Abrams Creek

48 35. Schematic of a 30 x 40-meter sampling plot

48 36. Diagram of the relationship of three 30 x 40-meter fixed sampling plots

49 37. Photo of stream sampling at Balsam Mountain

50 38. Photo of checking identification and recording data during stream sampling at 

Balsam Mountain

51 39. Schematic of a combined transect/coverboard survey design

53 40. Photo of coverboards

55 41. Photo of PVC pipes on pine trees in Okefenokee National Wildlife Refuge

56 42. Schematic of a survey design using paired PVC pipes located at 10-meter 

intervals around a pond’s perimeter

57

43-46. Photos of:

43. Leaf litterbag in Little River

58 44. Checking leaf litterbag at Little River

59 45. Recording data in field as storm approaches at Cataloochee Divide

68 46. Washing boots and stump ripper in bleach solution for biosecurity

72 47. Flow chart of: Decision path for helping design an amphibian monitoring 

program at Great Smoky Mountains National Park based on three levels of 

funding

Tables

12 1. Identification and life history of the nonpermanently aquatic salamanders of 

Great Smoky Mountains National Park

24 2. Identification and reproductive life history of the frogs of Great Smoky 

Mountains National Park
1

Monitoring Amphibians in 

Great Smoky Mountains 

National Park

By C. Kenneth Dodd, Jr.

— Abstract —

Amphibian species have inexplicably declined 

or disappeared in many regions of the world, and in 

some instances, serious malformations have been 

observed. In the United States, amphibian declines 

frequently have occurred even in protected areas. 

Causes for the declines and malformations probably 

are varied and may not even be related. The seemingly sudden declines in widely separated areas, however, suggests a need to monitor amphibian 

populations as well as identify the causes when 

declines or malformations are discovered. 

In 2000, the President of the United States and 

Congress directed Department of the Interior (DOI) 

agencies to develop a plan to monitor the trends in 

amphibian populations on DOI lands and to conduct 

research into possible causes of declines. The DOI 

has stewardship responsibilities over vast land holdings in the United States, much of it occupied by, or 

potential habitat for, amphibians. The U.S. Geological Survey (USGS) was given lead responsibility for 

planning and organizing this program, named the 

Amphibian Research and Monitoring Initiative 

(ARMI). Authorization carried the mandate to set up 

a national amphibian monitoring program on Federal 

lands, to develop the sampling techniques and biometrical analyses necessary to determine status and 

trends, and to identify possible causes of amphibian 

declines and malformations.

The biological importance of Great Smoky 

Mountains National Park has been recognized by its 

designation as an International Biosphere Reserve. 

As such, it is clearly the leading region of significance for amphibian research. Although no other 

region shares the wealth of amphibians as found in 

the Great Smokies (31 species of salamanders, and 

13 of frogs), the entire southern and mid-section of 

the Appalachian Mountain chain is characterized by 

a high diversity of amphibians, and inventories and 

monitoring protocols developed in the Smokies likely 

will be applicable to other Appalachian National Park 

Service properties.

From 1998 to 2001, USGS biologists carried 

out a pilot inventory and monitoring research project 

in Great Smoky Mountains National Park. A variety 

of inventory, sampling, and monitoring techniques 

were employed and tested. These included wide-scale 

visual encounter surveys of amphibians at terrestrial 

and aquatic sites, intensive monitoring of selected 
2

plots, randomly placed small-grid plot sampling, litterbag sampling in streams, monitoring 

nesting females of selected species, call surveys, and monitoring specialized habitats, such 

as caves. Coupled with information derived 

from amphibian surveys on Federal lands using 

various other techniques (automated frog call 

data loggers, PVC pipes, drift fences, terrestrial 

and aquatic traps), an amphibian monitoring 

program was designed to best meet the needs of 

biologists and natural resource managers after 

taking into consideration the logistics, terrain, 

and life histories of the species found within 

Great Smoky Mountains National Park. 

This report provides an overview of the 

Park’s amphibians, the factors affecting their 

distribution, a review of important areas of 

biodiversity, and a summary of amphibian life 

history in the Southern Appalachians. In addition, survey techniques are described as well as 

examples of how the techniques are set up, a 

critique of what the results tell the observer, and 

a discussion of the limitations of the techniques 

and the data. The report reviews considerations 

for site selection, outlines steps for biosecurity 

and for processing diseased or dying animals, 

and provides resource managers with a decision 

tree on how to monitor the Park’s amphibians 

based on different levels of available resources. 

It concludes with an extensive list of references 

for inventorying and monitoring amphibians. 

USGS and Great Smoky Mountains National 

Park biologists need to establish cooperative 

efforts and training to ensure that congressionally mandated amphibian surveys are performed in a statistically rigorous and 

biologically meaningful manner, and that 

amphibian populations on Federal lands are 

monitored to ensure their long-term survival. 

The research detailed in this report will aid 

these cooperative efforts.

The Florida Caribbean Science Center 

(now Florida Integrated Science Center) 

received funding in 1997 from the U.S. Geological Survey (USGS) Inventory and Monitoring 

(I&M) Program to conduct a pilot project for 

amphibians in the southeastern United States. 

After considering several locations, Great 

Smoky Mountains National Park (fig. 1) was 

selected for the survey because of its amphibian 

diversity and the large number of potential 

threats to its varied ecosystems (Brown, 2000). 

During the course of the next 4 years, a field 

research team of enthusiastic young biologists 

was assembled to collect information on the 

species richness and distribution of the Park=s 

amphibians. Researchers used a variety of sampling techniques, including 10 x 10-meter survey plots, “permanent” 30 x 40-meter plots, 

coverboards, litter-bag surveys, and a great 

number of time-constrained litter and stream 

searches. The team looked for previously 

reported rare species, sampled historic locations, investigated unique habitats (such as 

caves), and examined museum records and published literature. Survey activities and techniques were designed to optimize the use of 

available personnel within budget and logistic 

constraints. Survey teams sampled more than 

500 sites (fig. 2) and recorded data on more than 

10,000 amphibians. All parts of the Park were 

visited in all seasons and in all weather 

conditions.

The objectives of the Great Smoky 

Mountains National Park I&M program were 

to: (1) provide a geographically referenced 

inventory of the amphibian resources of the 

Great Smoky Mountains National Park; 

(2) provide indices of abundance of Park 

selected amphibian species, referenced to locations and habitat types; (3) develop and transfer 

to the Great Smoky Mountains National Park 

and National Park Service a series of protocols 

suitable for long-term monitoring of amphibian 

populations in the “Smokies” and other Appalachian parks; (4) evaluate current distributions 

and abundance of amphibian species as possible 

in the Park with literature reports of past investigations. This manual fulfills the third objective 

of the I&M program. Additional information on 

amphibian natural history, distribution, landscape ecology, trends analysis, and protocol 

development are published in Dodd and others, 

(2001), Waldron and others, (2003); Dodd, 

(2004), or is under development.

Introduction
3

84° 83°45′ 83°30′ 83°15′ 

35°45′35°30′

0 5 10 15 MILES0 5 10 15 KILOMETERS

Cades

Cove

Fontana Reservoir

Sugarlan

M d ountain

Middle

Prong

Little River

Abrams

Creek

Deep

Creek

Oconaluftee

River

Creek

Hazel Indian

Creek

Abrams Ranger Station and

Campground

TENNESSEE

NORTH CAROLINA Clingmans Dome

Sugarlands Visitor Center

Mount

Le Conte

Smokemont

Metcalf

Bottoms

Cherokee Indian

Reservation

Creek

Cosby

C

K

Cosby

Cosby Campground

Cades Cove

Campground

W.

Prong Littl

v

e Pigeon Ri er

Little Pigeon River

19

19

411

Silva

411

411

28

129 28

Foothills Parkway

73

129 411

Maryville

73

411

23

Bl

P

ue d Ri ge a

321

321 321

19

74

Pigeon Fork

Bryson City

Gatlinburg

NC

SC

GA

FL

MS AL

LA

TN

Great Smoky Mountains

National Park

GREAT SMOKY MOUNTAINS

NATIONAL PARK

EXPLANATION

Shaded relief indicates area of Great Smoky

Mountains National Park

Cherokee Indian Reservation

Cane

Creek

PSP

Whiteoak

Sink

Big Spring Cove

Big Cove

Base from U.S. Geological Survey digital data, datum nad83Albers Equal-Area Conic Projection,Standard parallels 29 30 and 45 30 , central meridian -83 00 °° ° ′′ ′

Figure 1. Great Smoky Mountains National Park, North Carolina and Tennessee.
4

84° 83°45′ 83°30′ 83°15′ 83°

35°45′

35°30′

0 5 10 MILES

0 5 10 KILOMETERS

TENNESSEE

NORTH CAROLINA

EXPLANATION

Great Smoky Mountains

National Park

Cherokee Indian

Reservation

Primary road (paved)

Secondary road (gravel)

Park trails

Park and Reservation

boundaries

Sampling sites, 1998-2001

Base from U.S. Geological Survey digital data, datum nad83,

Albers Equal-Area Conic Projection,

Standard parallels 29 30 and 45 30 ,

central meridian -83 00

° °

°

′ ′

′

Figure 2. Location of U.S. Geological Survey sampling sites in Great Smoky Mountains National Park, 1998 to 2001.

This manual is the result of collaborative survey 

efforts during 4 years of hard field work between 1998 

and 2001 in the Great Smoky Mountains. The author 

would like to thank the following individuals for their 

support, assistance, dedication, and companionship:

For their field collecting skill, expertise, and general good humor during all kinds of weather conditions, the USGS field crew from 1998 to 2001 (in 

alphabetical order): Jamie Barichivich, Jeff Corser, 

Elizabeth Domingue, Marian Griffey, Kelly Irwin, 

Christy Morgan, Rick >Bubba= Owen, Kevin G. 

Smith, and Jayme Waldron. For figure 7: Todd 

Campbell. For advice: James Petranka and Chuck 

Smith.

For their support and assistance: the National Park 

Service personnel at Great Smoky Mountains 

National Park, particularly Keith Langdon, Dana 

Soehn, and Jack Piepenbring.

For providing funding: Norita Chaney (USGS 

Inventory and Monitoring Program, Reston, 

Virginia), Russell Hall (Florida Integrated Science 

Center), and the USGS Amphibian Research and 

Monitoring Initiative (ARMI).

For providing research permits: National Park 

Service (Great Smoky Mountains National Park), 

North Carolina Wildlife Resources Commission, 

Tennessee Wildlife Resources Agency.

For providing the beautiful illustrations of larval 

amphibians: Jacqualine Grant.

For reading the manuscript and making valuable 

comments: Jamie Barichivich, Marian Griffey, 

Russell Hall, Steve Johnson, Keith Langdon, and 

Jennifer Staiger.

Acknowledgments
5

programs are not considered in detail. However, 

references are provided at the end of this guide 

(see References on Inventorying and Monitoring Amphibians).

Future amphibian monitoring within 

Great Smoky Mountains National Park will be 

linked to the U.S. Department of Interior (DOI) 

Amphibian Research and Monitoring Initiative 

(ARMI). Standardized methods of data collection, entry, and analysis currently are being 

developed by ARMI researchers for all DOI 

lands. Pertinent information will be made available to Federal agencies and ARMI partners 

through ARMI=s web site: 

edc2.usgs.gov/armi/

A cautionary note: There is always the 

danger that site information will be misused 

by criminal elements to find amphibians in 

order to collect them. This is true in National 

Parks and on other Federal lands, as well as on 

private lands. None of the amphibians found 

in Great Smoky Mountains National Park are 

endangered or threatened under the Federal 

Endangered Species Act of 1973, as amended, 

although several species, such as Hellbenders, 

are protected by state law. Locations of many 

of the Park=s amphibians, including its 

endemic salamanders, are well known via the 

published scientific literature and on records 

attached to museum specimens. Therefore, it 

seems unlikely that mentioning Park locations 

in this guide will increase the probability of 

collection, especially when these species are 

found readily, and often in greater abundance, 

outside the Park. For example, the Mole Salamander, Southern Zigzag Salamander, and 

Mud Salamander might be considered Arare@

or Aisolated@ within Great Smoky Mountains 

National Park, yet very large and widespread 

populations of these species are found in the 

Tennessee Valley and elsewhere. Still, Park 

Service employees and research scientists 

working within the Park, including field survey teams, must be observant for illegal collectors and immediately report suspicious 

activities to law enforcement personnel.

HOW TO USE THIS GUIDE

Monitoring Amphibians in Great Smoky 

Mountains National Park is meant to help 

National Park Service natural resource biologists, university researchers, nongovernmental 

biologists, and the interested public understand 

and overcome some of the biological and nonbiological constraints to setting up a large-scale 

inventory and monitoring program for amphibians inhabiting the Great Smoky Mountains. 

Some of the information applies only to 

amphibians within the Great Smokies, whereas 

information on setting up inventory and monitoring programs may have more broad applications with regard to Appalachian amphibians. 

Many persons who use this guide will be familiar with basic amphibian biology, but others will 

require a refresher course or will be unfamiliar 

with amphibian life histories.

This guide serves as a companion volume 

to Dodd (2004) and, for that reason, information 

in that work has not been duplicated except 

when absolutely necessary. There is usually an 

exception to every generalization discussed 

below, and biologists should expect to encounter species outside of their “normal” habitat, 

that often do not fit identification information, 

or that have unusual behavioral patterns. Extensive information is not included on threats to 

amphibians (for example, habitat loss and alteration, disease, nonindigenous species, climate 

change, toxic chemicals, UVB, malformations) 

or the various reasons why amphibians are vulnerable to environmental problems (including 

their biphasic life cycle, skin permeability, and 

the complex morphological and biochemical 

transformations which accompany metamorphosis). These topics are dealt with in more 

detail elsewhere (Dodd, 1997, 2004; Alford and 

Richards, 1999; Corn, 2000; Houlahan and 

others, 2000).

All of the potential sampling protocols, 

techniques, and methods of data analysis that 

may accompany, or be required for, a largescale amphibian inventory and monitoring 

program cannot be discussed within one short 

guide. For this reason, many specialized techniques are not discussed, instructions are not 

provided for making traps, and statistical 
6

AMPHIBIANS OF THE GREAT SMOKY MOUNTAINS

Species Richness

A total of 31 salamanders and 13 frogs have been recorded from the Great Smoky Mountains 

National Park. Note that common names are capitalized, and that species names (consisting of a 

genus and specific epithet) are italicized. Species codes allow data to be entered in shorthand format. To minimize data entry errors, species codes should be either all capitalized or all in lower 

case letters. Capitals and lower-case letters should not be intermixed. Using accepted and standardized common and scientific names (Crother, 2000), the amphibians are:

Common name Scientific name Suggested species

code

Salamanders

Spotted Salamander Ambystoma maculatum AMA

Marbled Salamander Ambystoma opacum AOP

Mole Salamander Ambystoma talpoideum ATA

Green Salamander Aneides aeneus AAE

Hellbender Cryptobranchus alleganiensis CAL

Seepage Salamander Desmognathus aeneus DAE

Spotted Dusky Salamander Desmognathus conanti DCO

Imitator Salamander Desmognathus imitator DIM

Shovel-nosed Salamander Desmognathus marmoratus DMA

Seal Salamander Desmognathus monticola DMO

Ocoee Salamander Desmognathus ocoee DOC

Black-bellied Salamander Desmognathus quadramaculatus DQU

Santeetlah Salamander Desmognathus santeetlah DSA

Pigmy Salamander Desmognathus wrighti DWR

Three-lined Salamander Eurycea guttolineata EGU

Junaluska Salamander Eurycea junaluska EJU

Long-tailed Salamander Eurycea longicauda ELO

Cave Salamander Eurycea lucifuga ELU

Blue Ridge Two-lined Salamander Eurycea wilderae EWI

Spring Salamander Gyrinophilus porphyriticus GPO

Four-toed Salamander Hemidactylium scutatum HSC

Common Mudpuppy Necturus maculosus NMA

Eastern Red-spotted Newt Notophthalmus viridescens NVI

Northern Slimy Salamander Plethodon glutinosus PGL

Jordan=s Salamander Plethodon jordani PJO

Southern Gray-cheeked Salamander Plethodon metcalfi PME

Southern Appalachian Salamander Plethodon oconaluftee POC

Southern Red-backed Salamander Plethodon serratus PSE

Southern Zigzag Salamander Plethodon ventralis PVE

Mud Salamander Pseudotriton montanus PMO

Black-chinned Red Salamander Pseudotriton ruber PRU

Frogs

Northern Cricket Frog Acris crepitans ACR

American Toad Bufo americanus BAM

Fowler=s Toad Bufo fowleri BFO

Eastern Narrow-mouthed Toad Gastrophryne carolinensis GCA

Cope=s Gray Treefrog Hyla chrysoscelis HCH

Spring Peeper Pseudacris crucifer PCR

Upland Chorus Frog Pseudacris feriarum PFE

American Bullfrog Rana catesbeiana RCA

Northern Green Frog Rana clamitans RCL

Pickerel Frog Rana palustris RPA

Northern Leopard Frog Rana pipiens RPI

Wood Frog Rana sylvatica RSL

Eastern Spadefoot Scaphiopus holbrooki SHO
7

 Amphibian taxonomy and systematics 

within the southern Appalachians are topics of 

intense debate among biologists. Rationale for 

using the listed names is provided by Dodd 

(2004).

Habitats and Distribution

Five major forest communities are recognized within the Great Smoky Mountains 

National Park, although 80 percent of the Park 

falls within the Eastern Deciduous Forest Ecosystem (Houk, 1993). Some botanists have further subdivided the vegetation into as many as 

67 florally distinct communities. No one species of amphibian is associated entirely with a 

single forest community, although some of the 

high-elevation salamanders (Plethodon jordani, 

Desmognathus ocoee, D. wrighti) are more 

often found in the spruce-fir community than in 

other community types. Habitat structure, particularly one that retains moisture and high 

humidity, is important in shaping salamander 

distribution. The high-elevation coniferous forest appears ideal in providing shade, cover (in 

the form of coarse woody debris), and abundant 

surfaces for moisture condensation.

Figure 3. 

Spruce-fir forest 

at Indian Gap.

The spruce-fir forest (fig. 3) is dominated by Red Spruce (Picea rubens) and Fraser 

Fir (Abies fraseri), and is found generally above 

1,676 m (5,500 ft), although the community 

descends to 1,372 m (4,500 ft) in some locations 

and individual Red Spruce are found at even 

lower elevations. This is the Canadian Zone 

boreal forest of high moisture, cool or cold temperatures, and high humidity (Houk, 1993). 

Ground surface is often dense with fallen tree 

branches and trunks, and carpeted by thick layers of tree needles. Wet, rotten, woody debris 

and dense needle mats provide ideal hiding 

places for terrestrial salamanders. Streams originate in this habitat, usually beginning as small 

seeps and springs. As streams trickle through 

Five major forest communities are recognized within the Great Smoky Mountains 

National Park....
8

the dark-green forest, they gather momentum. 

Even at higher elevations, aquatic salamanders, 

particularly duskies (Desmognathus) and Blue 

Ridge Two-lined Salamanders (Eurycea 

wilderae), may be plentiful within the headwater streams.

At somewhat lower elevations (1,067-

1,524 m; 3,500-5,000 ft), deciduous northern 

hardwoods (fig. 4) predominate, such as Sugar 

Maples (Acer saccharum), American Beech 

(Fagus grandifolia), and Yellow Birch (Betula 

alleghaniensis). Many terrestrial and aquatic 

salamanders reach their lower or upper distributional range within this community; frogs are 

scarce. Cove hardwoods, the third community, 

comprise the most diverse forest community in 

the Smokies, one that is endemic to the southern 

Appalachian Mountains. It occurs generally 

below 1,372 m (4,500 ft) in sheltered valleys, 

and is dominated by Tulip Poplar (Liriodendron 

tulipifera), Dogwood (Cornus florida), Red 

Maple (Acer rubrum), Sweetgum (Liquidamber 

styraciflua), White Basswood (Tilia americana

var. heterophylla), Yellow Buckeye (Aesculus 

flava), and Black Birch (Betula lenta). Both 

hardwood communities have complex understory vegetation, often with much coarse woody 

debris, which provides cover for terrestrial salamanders. The streams through these hardwood 

forests are rocky and fast paced, and salamanders are very common along streamsides 

and in the water.

Two somewhat specialized forest communities are found in the Smokies. The hemlock

community (fig. 5) is dominated by Eastern 

Hemlocks (Tsuga canadensis), commonly 

Figure 4. Deciduous forest at Lynn Hollow.
9

called “spruce-pines” by natives of the southern 

mountains, and is common between 1,067-

1,524 m (3,500-5,000 ft) in elevation. 

Hemlocks descend to much lower elevations 

along cold mountain stream valleys, and overlap considerably with both hardwood forests 

and the spruce-fir forest of the higher 

elevations. Hemlocks are massive with tall, 

Figure 5. Hemlock forest at Chinquapin Knob.

Hemlocks are massive 

with tall, straight trunks. 

When they fall, they provide excellent habitat for 

salamanders....

straight trunks. When they fall, they provide 

excellent habitat for salamanders, both in the 

rotting wood and under exfoliating bark (fig. 6).

The pine-oak forest (fig. 7) occupies the 

drier areas of the Park, particularly the area west 

of Cades Cove and at mid-elevations on the 

North Carolina side of the Park. This forest is 

dominated by Southern Red (Quercus falcata), 
10

Northern Red (Q. rubra), Scarlet (Q. coccinea), 

Black (Q. velutina), and Chestnut (Q. prinus) 

Oaks, and by Pitch (Pinus rigida), White (P. 

strobus), and Table Mountain (P. pungens) 

Pines. Soils are dry, as is the leaf litter. Prior to 

human intervention, this community burned frequently in the western regions of the Park, and 

a fire-adapted vegetation community resulted. 

Terrestrial salamanders are few, and usually 

found only during cool, wet times of the year. 

Aquatic-breeding salamanders and frogs are 

found along streamsides, where they likely 

remain close to water. The bottomlands along 

Cane Creek and Abrams Creek likely formed a 

corridor from the Tennessee Valley into Cades 

Cove. As a result, amphibian species richness is 

surprisingly high, particularly for frogs.

Amphibians are not uniformly distributed 

throughout the Great Smoky Mountains 

National Park. There are wide-ranging species, 

species restricted to specialized habitats, and 

species found in only one area of the Park. 

Figure 6. Coarse woody debris in Cove forest at Roaring Fork. Note the pink survey flags marking the 

position of transects.
11

Monitoring programs will need to take the 

distribution of species into account to optimize 

time and financial resources. A few generalizations can be made about amphibian distribution 

and habitats within the Park.

SALAMANDERS

Terrestrial salamanders (see Life 

History) include species that are: restricted in 

distribution in the Great Smokies; wide ranging 

but not common species; and wide ranging in 

higher or lower elevations, and generally common. Because they do not have larvae, they 

must be sampled where they carry out their 

entire life cycle, usually on the forest floor 

and under leaf litter and other debris.

Figure 7. Oak-pine forest. Photographer: Todd Campbell, University of Tampa

Amphibians are not 

uniformly distributed 

throughout the Great 

Smoky Mountains National 

Park.
12

Table 1. Identification and life history of the nonpermanently aquatic salamanders of Great Smoky Mountains National Park[mos, months; yr, year; mm, millimeter; TL, total length; SVL, snout-vent length; ~, approximately; <, less than; >, greater than]

Species Egg deposition Hatching Larval period Hatching size Size and time of Metamorphosis Spots on dorsum Dorsal pattern Belly pattern T

Ambystoma maculatum Jan. to late Mar. (mountains-late Feb. 

to early Mar.); 4-7 weeks incubation

April-May 2-4 mos 12-17 mm TL 29-32 mm SVL; 43-60 

mm TL (to 75 mm TL if overwinter); midJune to August

dull olive green, no conspicuous markings

white or light tai

mfinda

A. opacum Oct-Nov (in pond by 

Sept.); 9-15 days incubation, but must be flooded 1-2 days

winter 5-7 mos 10-14 mm TL ~33 mm SVL; 49-58 

mm TL; late March mid-June

blackish, drab; older larvae have mottling on body

throat stippled; scattered melanophores on lateral sides

doexto 

A. talpoideum Sept. to Mar. (winter) winter to 

early spring

3-4 mos, but variable

~10 mm SVL 32 to 50 mm SVL; 

May to Sept.

black and yellow blotches along midline of back

dark band on midline (poor in some specimens)

yebl

Desmognathus conanti

early May to early July, perhaps to midAugust; 45-60 days incubation

July to early fall

<1 yr 8-12 mm SVL; 

12-20 mm TL

9-12 mm SVL, to 20 mm SVL; July to early fall?

5-8 pairs of even or alternating spots or blotches

sides with dorsolateral stripe; dorsum variable

spcotai

D. imitator late spring to early 

summer?

D. marmoratus late spring to early 

summer?; 10-12 weeks incubation

mid-Aug to mid-Sept.

3 yrs (10-20 mos)

11 mm SVL 26-38 mm SVL; May 

to Oct.

2 rows light spots dark, conspicuous 

light flecks on sides

sp

D. monticola mid-June to midAugust; 2 mos incubation early sum- mer to fall; Sept. 10-11 mos 11-12 mm SVL June-July 4-5 pairs light dor- sal spots between limbsD. ocoee July to early Aug. to 

Sept.; 52-74 days incubation

Aug to late Sept.

9-10 mos 13-18 mm TL 11-15 mm SVL; May 

to June

4-6 pairs of alternating light spots on dorsum

D. quadramaculatus May to June July to Sept. 3-4 yrs 11-16 mm SVL 35-42, to 54 mm SVL; mid-summer 6-8 pairs light spots between limbs light brown D. santeetlah early May to early 

July, perhaps to midAugust; 45-60 days incubation

July to early fall

<1 yr 8-12 mm SVL; 

12-20 mm TL

9-12 mm SVL, to 20 mm SVL; July to early fall?

4-5 pairs of even or alternating spots or blotches
13

Eurycea guttolineata

winter early to midMar? 3.5-5.5 mos (< 1 yr), but 

may overwinter

11-12.5 mm SVL

22-27 mm SVL, to 32 mm SVL; June to August

no paired light spots cream; uniformly 

stippled; then dark broad dorsolateral stripe; narrow middorsal stripe

immaculate do

nowaleg

E. junaluska at least by mid-May early June? 1-2 yr 7-9 mm SVL; 

11-13 mm TL

34-42 mm SVL; midMay to August deep olive green to brown no iridophores 

E. longicauda late autumn to early 

spring

Nov-March after 4-12 weeks

normally <1 yr (4-7 mos)

18-21 mm SVL; 40 mm TL;

23-28 mm SVL; > 50 mm TL if overwintering; mid-JuneJuly

cream colored; then uniformly dark, similar to adults; no paired spots

immaculate

E. lucifuga Sept. to Feb. 6-18 mos; 

most 12-15 mos

9-12 mm SVL; to 17.5 mm TL

31-37 mm SVL; to 70 mm TL; spring

sparse pigmentation with 3 longitudinal series of spots on the side

E. wilderae Feb. to May; 4-10 

weeks incubation

May to August

1-2 yr 7-9 mm SVL; 

11-14 mm TL

18-19 mm SVL in 1 yr, to 32 mm SVL in 2 yr; April to July

6-9 pairs light dorsolateral dusky light with iri- dophores 

Gyrinophilus porphyriticus

summer late summer 

to autumn

to 4 yr 18-22 mm TL 55-65 mm SVL, to 

70 mm high elevation; late June to August

light yellow brown to gray with fine flecking

Hemidactylium scutatum

Feb to May May-June? 21 to 61 

days

11-15 mm SVL; 17-25 mm TL; July?

nondescript, yellow brown; dorsal fin extends to head

Pseudotriton montanus

autumn to early winter

winter 15-17 mos 

to 29-30 mos

<13 mm SVL 35-44 mm SVL; midMay to Sept. light brown; older with widely scattered 

spots

immaculate 

P. ruber autumn to early winter; 3 mos incubation mid-Dec to mid-Feb 1.5 to 3.5 yr (27-31 mos) 11-14 mm TL 34-46 mm SVL; 62- 86 mm TL; May to 

July

light brown; weakly mottled or streaked

dull white 

Table 1. Identification and life history of the nonpermanently aquatic salamanders of Great Smoky Mountains National Park (Continued)

[mos, months; yr, year; mm, millimeter; TL, total length; SVL, snout-vent length; ~, approximately; <, less than; >, greater than]

Species Egg deposition Hatching Larval period Hatching size Size and time of Metamorphosis Spots on dorsum Dorsal pattern Belly pattern Ta
14

Monitoring programs can target each type of 

distributional pattern or habitat listed below, 

depending upon the objectives of the researchers and the funds and personnel available. For 

example, whereas a few people can easily monitor the status of the Southern Zigzag Salamander, a much more elaborate protocol will be 

necessary to monitor populations of the Southern Red-backed Salamander. A number of these 

species are syntopic, making multispecies monitoring a realistic objective. As much as possible, single species sampling and monitoring 

should be avoided in favor of multispecies 

sampling and data recording. Some examples of 

typical distribution patterns follow:

Species restricted in distribution

Southern Zigzag Salamander (Plethodon 

ventralis).

Wide ranging, but not common, species

Southern Appalachian Salamander (Plethodon oconaluftee).

Species that are common and wide

ranging at higher elevations

Pigmy Salamander (Desmognathus wrighti); 

Jordan=s Salamander (Plethodon jordani); 

Southern Gray-cheeked Salamander (Plethodon metcalfi).

Species that are common and wide

ranging at lower elevations

Northern Slimy Salamander (Plethodon glutinosus); Southern Red-backed Salamander 

(Plethodon serratus).

River-dwelling salamanders inhabit only 

the largest of the Smokies’ rivers (fig. 8), 

including Little River, Middle Prong, Oconaluftee River, Little Pigeon River, Abrams 

Creek, the lower reaches of Deep Creek and, 

perhaps, Hazel Creek. There are only two true 

river-dwelling salamanders in the Great Smokies, the Hellbender (Cryptobranchus alleganiensis), known presently only from Little 

River, Oconaluftee River, and Deep Creek 

Figure 8. Middle Prong at Tremont.
15

Larvae are found near the shore, and the adults 

inhabit streambanks for at least part of the year. 

However, this species also inhabits some 

smaller streams, and it is by no means a Ariverdwelling@ species.

Creek and stream salamanders have larvae that develop in the creeks and streams of the 

Park (figs. 10-12), whereas the adults may be 

(Nickerson and others, 2002; Dodd, 2004) 

(fig. 9), and the Common Mudpuppy (Necturus 

maculosus), known only from Little River and 

Abrams Creek. One additional salamander, the 

Junaluska Salamander (Eurycea junaluska), 

tends to be associated with some of the Park=s 

larger western and northwestern streams and 

rivers on the Tennessee side of the Smokies. 

Figure 9. Ideal 

habitat for 

Hellbenders in 

Lower Abrams 

Creek.

Figure 10. 

Small stream in 

unnamed 

tributary to Falls 

Branch.
16

Figure 11. Medium-sized stream in normal flow at Roaring 

Fork.

aquatic, semi-aquatic, or even terrestrial to a 

greater or lesser degree. Many of these species 

are widespread in the Park because of the large 

number of creeks and streams available for colonization. A few species are found only at 

higher mountain elevations (for example, the 

Ocoee and Imitator Salamanders), whereas others are lowland species (Spotted Dusky, Threelined, and Long-tailed Salamanders). Instead of 

a circumscribed area, their habitat is often linear, following the streams and streamsides. The 

dusky salamanders (Desmognathus) are prominent in this group, but there are many exceptions 

to each habitat categorization listed below. Even 

Black-bellied Salamanders have been found 

well above the forest floor in rock crevices 

among boulders at considerable distances from 

water. Monitoring adults and larvae of these 

species requires very different techniques, and 

may require sampling very different types of 

habitats.

Nearly aquatic species

Shovel-nosed Salamander (Desmognathus 

marmoratus).

Predominantly aquatic and streamside 

species

Spotted Dusky Salamander (Desmognathus 

conanti); Seal Salamander (Desmognathus 

monticola); Black-bellied Salamander 

(Desmognathus quadramaculatus); Santeetlah Salamander (Desmognathus santeetlah); 

Many of these species 

are widespread in the 

Park because of the large 

number of creeks and 

streams available for 

colonization.

Figure 12. Medium-sized stream 

in high water at Whiteoak Flats 

Branch.
17

Junaluska Salamander (Eurycea junaluska). 

The extremely high elevation areas where 

some streams first appear may be devoid of 

salamanders if the water emanates from the 

Anakeesta rock formation (Dodd, 2004).

Species with aquatic larvae but are largely 

terrestrial as adults

Imitator Salamander (Desmognathus imitator); Ocoee Salamander (Desmognathus 

ocoee); Three-lined Salamander (Eurycea 

guttolineata); Long-tailed Salamander 

(Eurycea longicauda); Blue Ridge Twolined Salamander (Eurycea wilderae).

A few salamanders require very specialized habitats in the Great Smokies, or at least 

are usually found in these habitats. Some of 

these species have larvae which are found in the 

same streams and creeks as the preceding species, although the adults prefer to leave the 

streams. Whereas the larvae may be relatively 

easy to survey, adults often can be quite difficult 

to find with any regularity. One species, the 

Seepage Salamander, does not have a larval 

stage, and the adults are only found in wet 

seeps.

Cave inhabitants (fig. 13)

Cave Salamander (Eurycea lucifuga).

 Known only from Stupkas Cave, the Calf 

caves, and one record from Whiteoak Sink. 

Other salamanders in the Smokies may live in 

caves, especially around the entrances (Dodd 

and others, 2001). The larvae of some salamanders (for example, E. longicauda in 

Gregorys Cave) develop in pools well inside 

cave passages (fig. 14).

Rock face inhabitants (fig. 15)

Spotted Dusky Salamander (Desmognathus 

conanti); Seal Salamander (Desmognathus 

monticola). 

Permanent to near permanent wet rock 

walls with hiding places, particularly along 

trails, road cuts, and in the vicinity of waterfalls, 

especially at lower elevations.

Figure 13. Entrance to Gregorys Cave.
18

Figure 15. Rock face near Double Gap.

Figure 14. Rimstone pools and cave pool at 

Gregorys Cave. Salamander larvae develop 

in the pools, although they are unlikely to 

complete metamorphosis.

A few salamanders require 

very specialized habitats in 

the Great Smokies, or at 

least are usually found in 

these habitats.
19

Spring Inhabitants

Spring Salamander (Gyrinophilus porphyriticus); Black-chinned Red Salamander 

(Pseudotriton ruber).

Inhabitants of swampy and mucky habitats 

(fig. 16)

Mud Salamander (Pseudotriton montanus).

Known only from a few scattered locations in the lowlands of the northern side of the 

Park.

Inhabitants of wet seepages (fig. 17)

Seepage Salamander (Desmognathus 

aeneus). 

Known only from drainages on the 

southwestern side of the Park.

Finally, there are salamanders that 

breed in ponds, and it is virtually only at this 

time that these species can be censused. Five 

species fall into this category: the Spotted 

Salamander (Ambystoma maculatum); Marbled Salamander (A. opacum); the rare Mole 

Salamander (A. talpoideum); Four-toed 

Salamander (Hemidactylium scutatum); and 

Eastern Red-spotted Newt (Notophthalmus 

viridescens). Breeding ponds are limited 

within the Park, being concentrated in Cades 

Cove and nearby Big Spring Cove (the four 

Finley-Cane sinkhole ponds), the Cane Creek 

drainage, and at scattered localities between 

Sugarlands and Cades Cove along Little River 

Figure 16. Former trout pond mucky habitat in 

Cataloochee.

(at the Sinks and ditches along the road to 

Tremont). These locations are on the Tennessee 

Figure 17. Seep at Big Spring Cove.

side of the Park. Although beaver 

ponds are found in Bone Valley 

and Big Cove in North Carolina, 

and small scattered ditches and 

wetlands occur in Cataloochee 

Valley, no pond salamanders are 

known to breed in them.

Mud Salamanders are known 

only from a few scattered locations in the lowlands of the 

northern side of the Park.
20

are used by frogs and toads: ponds (natural, as 

well as of beaver or human origin); woodland 

pools; grassy ditches, pools, and rivulets; and 

larger streams and rivers.

Ponds

Pond distribution is limited in the 

Smokies, being confined mostly to Cades Cove, 

Big Spring Cove, and two beaver ponds. The 

Figure 18. Gum 

Swamp at Cades 

Cove in high water.

Figure 19. Gum 

Swamp at Cades 

Cove when dry.

FROGS

Frogs in the Great Smoky Mountains 

National Park require water for breeding and 

for tadpole development. As such, the diversity 

and distribution of frogs are not as great in the 

mountains as in the adjacent lowlands of the 

Tennessee Valley and Atlantic Coastal Plain. In 

the Smokies, four major types of breeding sites 
21

most important frog-breeding ponds are Gum 

Swamp (figs. 18, 19), Gourley Pond (figs. 20, 

21), Methodist Church Pond (fig. 22), and the 

sewage-treatment pond (all in Cades Cove); the 

four sinkhole ponds in Big Spring Cove (also 

known as the Finley-Cane ponds); and the beaver ponds in Bone Valley and Big Cove 

(fig. 23). Species that commonly use these 

ponds are the American Toad (Bufo americanus), Cope=s Gray Treefrog (Hyla chrysoscelis), 

Figure 20. Gourley 

Pond at Cades Cove 

in high water.

Figure 21. Gourley 

Pond at Cades Cove 

when dry.

Northern Green Frog (Rana clamitans), 

Pickerel Frog (R. palustris), Wood Frog (R. 

sylvatica), and Eastern Spadefoot (Scaphiopus 

holbrooki), known only from Gum Swamp. 

American Bullfrogs (R. catesbeiana) also have 

been heard at the beaver pond in Big Cove. 

Some of these ponds dry completely as the 

summer progresses, particularly Gum Swamp 

(fig. 19), Gourley Pond (fig. 21), and the 

Finley-Cane sinkhole ponds.
22

Figure 22. Methodist Church Pond at Cades Cove.

Figure 23. Beaver pond at Big Cove.
23

Woodland Pools

Woodland pools are scattered at 

various areas within the Park. They 

range from a few centimeters deep to 

about 0.5 m, and they usually dry as 

summer progresses. Woodland pools are 

located in level ground at Cosby, Sugarlands, Metcalf Bottoms, Big Spring 

Cove, Little Cataloochee Valley, 

throughout the Cane Creek drainage 

(fig. 24), Cades Cove (especially along 

Abrams Creek at the western edge of the 

cove), and doubtless in other areas of the 

Park. Amphibians that use these small 

pools for breeding include the Eastern 

Red-spotted Newt (Notophthalmus 

viridescens), American Toad (Bufo 

americanus), Cope=s Gray Treefrog 

(Hyla chrysoscelis), Northern Green 

Frog (Rana clamitans), Pickerel Frog 

(R. palustris), and Wood Frog 

(R. sylvatica).

Grassy Ditches, Pools, and Rivulets

Grassy ditches, pools, and rivulets 

are generally shallow, open-canopied 

habitats, with a grassy vegetation where 

concealment and breeding sites are 

available (fig. 25). Only two places in 

the Park contain much of this habitat: 

Cades Cove and Cataloochee Valley. 

Frogs found here include the American 

Toad (B. americanus); Eastern Narrowmouthed Toad (Gastrophryne carolinensis), known only from grassy pools at 

the Abrams Creek Ranger Station and at 

Shields Pond in Cades Cove; Spring 

Peeper (Pseudacris crucifer); Upland 

Chorus Frog (Pseudacris feriarum); and 

the ubiquitous Wood Frog (R. sylvatica). These habitats normally dry rapidly with 

the warm weather, although the rivulets and 

some pools in Cades Cove may persist well into 

summer.

Streams and Rivers

A few species of frogs breed in the shallows of rivers and larger streams. In the Great 

Figure 24. Woodland drainage pool at Cane Creek.

Smokies, the American Bullfrog=s (R. catesbeiana) large tadpoles are conspicuous in Abrams 

Creek near the Abrams Creek Ranger Station. 

Additional species, such as Fowler=s Toad 

(B. fowleri), breed in the backwaters formed 

from flooding along streams and rivers. Other 

frogs, such as Northern Green Frogs (R. clamitans), are found along streambanks during the 

nonbreeding seasons.
24

Table 2. Identification and Life History of the Frogs of Great Smoky Mountains National Park

[<, less than; >, greater than; mm, millimeter; cm, centimeter; m, meter; m2, square meters]

Species Eggs Tadpole description Breeding 

times Larval period Metamorph size

Acris 

crepitans

eggs deposited singly; 1 gelatinous envelope, >2.3 mm in 

diameter; deposited in shallow 

water among stems of grass or 

on bottom; 250 eggs per complement

a medium-sized light to medium-gray tadpole; 

throat light; tail musculature mottled or reticulated; usually a very distinctive "black flag" on 

the tail tip; tail long and narrow; anus dextral (to 

the right); oral disk emarginate; most 30-36 mm 

total length, rarely to 46 mm

April to June, 

possibly into 

July

35-70 days, 

based on Acris 

crepitans blanchardi

10-15 mm

Bufo 

americanus

eggs in strings with gelatinous 

casings; 2 envelopes present; 

strings long, to 60 m; 15-17 

eggs per 25 mm; 4,000-12,000 

eggs on bottom of quiet pools

body round or oval in dorsal view; eyes dorsal 

(looks cross-eyed); nostrils large; color dark 

brown to black; dorsal portion of the body unicolored; venter with aggregate silvery or copper spots; snout sloping in lateral view; tail 

musculature distinctly bicolored; anus medial 

(in the center); spiracle is distinctly on left side 

of body

spring 

(March-April)

50-65 days 7-12 mm

B. fowleri eggs in strings with gelatinous 

casings; 1 envelope present and 

<5 mm in diameter; strings 2.4-

3 m with 17-25 eggs per 25 

mm; 5,000-10,000 eggs; in tangled mass around vegetation

body round or oval in dorsal view; eyes dorsal 

(looks cross-eyed); nostrils large; color dark; 

dorsal portion of body slightly mottled; snout 

rounded in lateral view; tail musculature often 

not distinctly bicolored; anus medial (in the 

center); spiracle is distinctly on left side of 

body

April to July 40-60 days 7.5-11.5 mm

Gastrophryne 

carolinensis

eggs in small surface film that 

has a mosaic structure; envelope a truncated sphere; mass 

round or square; 10-150 eggs 

per mass; in any depression 

with water, but not deep pools

a small jet-black tadpole with lateral white to 

pink stripes on posterior portion of body 

extending to the tail musculature. Viewed from 

the side, the head comes to a point; body round 

in dorsal view; eyes wide set and lateral; anus 

median; jaws do not have keratinized sheaths, 

and the oral disc and labial teeth are absent

mid-May to 

mid-August

20-70 days 8.5-12 mm

Hyla 

chrysoscelis

eggs in small surface film, but 

envelope not in truncated 

sphere; no mosaic structure; 5-

40 eggs per mass; in shallow 

ponds attached loosely to vegetation, or free. Air bubbles 

present.

small to medium-sized grayish tadpole with a 

high dorsal tail fin; dorsal tail fin height equal 

to or greater than musculature height; tail long, 

with black blotches; background color of 

mature tail orange to scarlet; throat rarely pigmented; dorsal fin never extends anterior to 

midway between the spiracle and eye; anus 

dextral (to the right); oral disk not emarginate

April to June, 

but calls occasionally heard 

at other times 

of the year

45-65 days 13-20 mm

Pseudacris 

crucifer

eggs deposited singly in shallow water near bottom among 

vegetation; one gelatinous envelope.

a small-sized deep-bodied tadpole with a 

medium-sized tail; tail musculature mottled; 

fins clear or with blotches; no dots on body; 

snout square when viewed dorsally; anus dextral (to the right); oral disk not emarginate

late winter to 

early spring 

(February to 

April); calls 

occasionally 

heard at other 

times of the 

year

90-100 days 9-14 mm

P. feriarum egg mass in lump, but loose 

irregular cluster; 1 envelope, 

3.6-4.0 mm; deposited in 

marshy areas and pools in matted vegetation

small olive to black tadpole with a bronze 

belly; tail medium; anus dextral (to the right); 

oral disk not emarginate; tadpoles develop 

rapidly

February to 

April.

50-60 days 8-12 mm
25

Rana 

catesbeiana

eggs in large surface film in 

form of a disc; 10,000-12,000 

eggs per disc; deposited among 

water plants or brush; 1 gelatinous envelope

large olive to grayish green tadpole with small 

widely spaced small spots (dots) covering the 

body and tail; venter straw; eyes bronze; body 

oval and round in dorsal view; eyes dorsal or 

dorsolateral; nostrils small compared with 

eyes; lower jaw wide; anus dextral (to the 

right); oral disk emarginate

late spring 

and throughout the summer. Calls 

may be heard 

at other times 

of the year

1-2 years 31-59 mm

R. clamitans eggs in surface film; mass 

<0.09 m2

; 1,000-5,000 per 

mass; attached to vegetation or 

free; 2 gelatinous envelopes

large (but not deep bodied) olive green tadpole 

with large dark spots, generally with a white 

throat; belly deep cream without iridescence; 

body oval and round in dorsal view; eyes dorsal or dorsolateral; nostrils small compared 

with eyes; tail green mottled with brown; 

lower jaw wide; anus dextral (to the right); oral 

disk emarginate

late April to 

late July or 

even early 

August. Calls 

may be heard 

at other times 

of the year

to 1 year 23-38 mm

R. palustris eggs in firm regular cluster; 

brown above and yellow below; 

mass a sphere 38-100 mm in 

diameter; 2 envelopes present; 

2,000-4,000 eggs; mass deposited 75-100 mm to 91 cm under 

water; attached to debris and 

vegetation

large, full, deep-bodied tadpole; olive green 

shading through yellow on sides; venter cream, 

back marked with fine black and yellow spots; 

belly with blotches of white; venter iridescent, 

viscera visible; tail very dark, black blotches 

can aggregate to purple-black; body oval and 

round in dorsal view; eyes dorsal or dorsolateral; nostrils small compared with eyes; lower 

jaw narrow; anus dextral (to the right); oral 

disk emarginate

late winter to 

spring (midMarch-April)

70-80 days 19-27 mm

R. pipiens mass a firm regular cluster; 

3,500-6,500 eggs close together 

in mass; 2 envelopes present; 

outer envelope 5 mm; eggs 

black above and white below; 

deposited near surface, usually 

attached to grasses and vegetation, sometimes free

large, deep-bodied tadpole; dorsally dark 

brown, covered with small gold spots; belly 

deep cream, with bronze iridescence; viscera 

visible; throat translucent and more extensive 

than Pickerel Frog; similar in appearance to 

Green Frog, but darker; body oval and round in 

dorsal view; eyes dorsal or dorsolateral; nostrils small compared with eyes; lower jaw narrow; anus dextral (to the right); oral disk 

emarginate

probably early 

March to 

early May

60-80 days 18-31 mm

R. sylvatica eggs in firm regular cluster; 

black above and white below; 

mass a sphere 38-100 mm in 

diameter; 2 envelopes present; 

2,000-4,000 eggs; mass deposited 75-100 mm to 91 cm under 

water; attached to debris and 

vegetation

medium-sized tadpole with usually very dark 

to gray coloration, and with a faint light stripe 

of cream, white or gold along the upper jaw 

(like a mustache); venter cream with belly 

slightly pigmented at sides; body oval and 

round in dorsal view; eyes dorsal or dorsolateral; nostrils small compared with eyes; anus 

dextral (to the right); oral disk emarginate; tail 

quite long; dorsal crest high extending on to 

body

winter and 

early spring 

(mid-December to March)

45-85 days 16-18 mm

Scaphiopus 

holbrooki

eggs in loose irregular cylinder 

or band; mass 25-75 mm wide 

and 25-305 mm long; deposited 

on stems of plants/grass; 

1 gelatinous envelope; 200 per 

packet

a small dark tadpole, bronze to brown with 

close-set tiny orange spots; body round or oval 

in dorsal view; eyes close-set and dorsal, iris 

black; head wide relative to body width; tail 

short, with tip blunt and rounded; anus medial 

(in the center); spiracle is ventrolateral. Often 

found in "schools" of hundreds of tadpoles

only heard 

calling once 

(July 12, 

1999). Probably any time 

from March to 

October

14-60 days 8.5-12 mm

Table 2. Identification and Life History of the Frogs of Great Smoky Mountains National Park (Continued)

[<, less than; >, greater than; mm, millimeter; cm, centimeter; m, meter; m2, square meters]

Species Eggs Tadpole description Breeding 

times Larval period Metamorph size
26

Other Breeding Sites

Four minor types of wetlands and aquatic 

sites are used occasionally by frogs for breeding 

in the Great Smokies. American toads (B. americanus) breed in the backwaters along the north 

shore of Fontana Reservoir, although reservoirs (fig. 26) are generally depauperate of 

amphibians. Small, usually closed-canopied, 

swampy and mucky wetlands (for example, 

those found along Indian Creek, at Smokemont, 

and at the old trout pond in Cataloochee; see 

fig. 16) are used by Wood Frogs (R. sylvatica). 

Wood Frogs are quite variable in their choice of 

breeding sites, even to depositing eggs in 

human-enlarged spring pools and roadside 

ditches. Indeed, virtually any pool in late winter 

to early spring is likely to be colonized by 

breeding Wood Frogs.

Life History

Terrestrial Salamanders (Plethodontidae). The life cycle of terrestrial plethodontids 

takes place in a multidimensional space. Naturalists tend to think of salamanders as surfacedwelling, but surface activity is only a small 

part of the life cycle of a terrestrial salamander. 

Most terrestrial species probably do not have a 

very large home range on the ground surface, 

including beneath debris and litter. They spend 

a considerable part of their lives underground, 

and biologists really know very little about their 

life history, especially their time spent underground and the depth and range of underground 

lateral movement. In addition, terrestrial species occasionally become arboreal during the 

night or under rainy conditions; salamanders 

often take refuge under loose bark. Salamanders 

at different life stages may remain nearly 

Figure 25. Grassy pool at Cades Cove.
27

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, detection probabilities may change with life stage within a habitat. 

The eggs of some terrestrial species have never 

been seen, and nests have been located only 

with extreme infrequency. Some plethodontids 

may be long-lived (5-10 years).

Semi-Aquatic Salamanders (Ambystomatidae, Plethodontidae, Salamandridae). All 

attributes that apply to terrestrial salamanders 

apply to semi-aquatic salamanders in terms of 

surface and underground habitat use. 

Semi-aquatic salamanders, however, require 

water for reproduction. For mole salamanders 

(Ambystoma) and newts (Notophthalmus), 

breeding sites are usually standing water 

(ponds, ditches) free of fishes. For semi-aquatic 

plethodontids, breeding sites include seeps and 

streams from little trickle trails to sizeable 

streams or rivers. Adults (mole salamanders and 

newts) may migrate synchronously to breeding 

sites 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 censused, but herpetologists 

know little about what proportion of a population breeds annually, and from what area they 

are drawn. Males and females may not stay for 

equal amounts of time during the breeding season, even when the breeding season is extended.

Stream-breeding species may live permanently in the streams (Desmognathus marmoratus), streamsides (many other Desmognathus), 

or at various distances from the stream (D. imitator, Gyrinophilus, Pseudotriton). Distances 

may range from a few meters to hundreds of 

Figure 26. Chilhowee Lake at mouth of Abrams Creek.
28

meters away, and breeding migrations are not 

synchronized. Little is known about spatial distribution during terrestrial nonbreeding times. 

For some species (for example, Hemidactylium 

scutatum) virtually nothing is known about their 

lives away from woodland pools and streams/

ditches outside of the breeding season. For certain species (D. quadramaculatus) adults can be 

censused streamside, whereas adults of other 

species (D. imitator) can be readily found in terrestrial habitats; some species (Pseudotriton) 

can be found terrestrially as adults usually only 

by luck, and the adults of a few species 

(egg-brooding adult female Hemidactylium) are 

observed only during the breeding season.

All eggs of semi-aquatic salamanders are 

deposited in water, and the egg masses of some 

species (Ambystoma) can be censused easily. All 

semi-aquatic species have 

larvae which remain in a 

larval stage from a few 

months to as long as 2-3 years. 

Paedomorphosis (the ability to 

breed while maintaining a larval appearance) occurs in a few 

species (Ambystoma 

talpoideum) under favorable conditions, but no salamanders from 

the Park are known to be paedomorphic. Larvae metamorphose and presumably take up adult habits, but nothing is 

known concerning dispersion for most species. 

Maturation can range from one to many years, 

depending on species. Individuals of some species (Ambystoma, Notophthalmus, large Desmognathus) may live 10-15 or more years.

Aquatic salamanders (Cryptobranchidae, 

Proteidae). Little is known about the life history 

of most of these species, except for Cryptobranchus. Species within these families are entirely 

aquatic. The spatial use of habitat is largely 

unstudied except for Hellbenders, which are 

known to have home ranges and to guard nesting sites. Fully aquatic species (Cryptobranchus, Necturus) inhabit medium to large 

streams and rivers in the southern Appalachians. Hellbenders may live 25 or more years. 

Nothing, however, is known about longevity of 

the Common Mudpuppy (Proteidae: Necturus), 

because the larvae are little known and, for the 

most part, rarely seen.

Frogs. All of the frogs in the southern 

Appalachians have a “typical” amphibian life 

cycle. Adults move to a breeding site, deposit 

eggs that hatch into larvae (tadpoles), metamorphose to juveniles, disperse, and grow until they 

are ready to repeat the cycle. For most species, 

however, many questions about the life cycle 

remain unanswered (what percentage is breeding in any one year, where do juveniles go, how 

far do adults disperse). Larval periods may be 

extremely brief (days in Scaphiopus) to 

extremely long (years in some Rana). Breeding 

may be synchronous (spadefoots, many ranids) 

or extended (Rana catesbeiana). Even when 

synchronous and explosive (Rana sylvatica), 

the actual breeding date may extend over a 

period of months (December to 

March) as adults wait for the 

right combination of environmental conditions. Adults 

(and perhaps juveniles) of 

many frog species spend 

most of their lives away 

from the breeding sites. 

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 to sample them. However, the terrestrial sites are extremely important to survival 

since individuals spend most of their lives as 

terrestrial predators.

Although most species of frogs call during the breeding season, some species do not or 

they have only weak voices that do not carry far. 

Calling times are variable among species; some 

call during the day, some call at dusk and during 

the early evening, and some call only between 

midnight and early dawn. Some species call 

only during rains, whereas others will call most 

evenings of the breeding season. Some frogs 

breed in winter (even in the mountains of the 

South), others breed in the spring or summer, 

whereas others call during an extended breeding 

season. Calling times and seasons also vary latitudinally and perhaps with elevation.
29

Areas of Particular Amphibian Species 

Richness

Three areas within Great Smoky Mountains National Park are particularly rich in 

amphibians. Two (Cades Cove, Cane Creek 

drainage) are lowland sites, whereas the third is 

the high-elevation spruce-fir forest. The lowland sites are similar in amphibian species composition; they are rich in species because they 

are the only two sizeable lowland areas within 

the Park with a large variety of wetlands. As 

such, they contain most of the frogs and pondbreeding salamanders. Both areas share species 

affinities with the herpetofauna of the Tennessee Valley, from whence lowland amphibians 

colonized Cane Creek and Cades Cove (via 

Abrams Creek). On the other hand, the highelevation amphibians are composed entirely of 

salamanders, and two species (Plethodon jordani, Desmognathus imitator) are virtually 

endemic to the Park (D. imitator is found also in 

the Plott Balsams). Other high-elevation species 

in the spruce-fir forest (for example, D. ocoee, 

D. wrighti, P. metcalfi) are found in other 

restricted regions of the Southern Appalachians. 

These three areas should be the special focus of 

amphibian monitoring activities.

Identification

Most biologists working at Great Smoky 

Mountains National Park should be able to 

identify the majority of the amphibians that they 

observe by using a combination of the color 

photographs, species descriptions, and identification/life history tables found in this manual 

and in Dodd (2004). Some individual animals 

may be impossible to identify with certainty. 

Larvae, especially small salamander larvae and 

tadpoles, often cannot be distinguished without 

microscopic examination. Adult salamanders, 

especially the duskies (Desmognathus), are 

notoriously variable with overlapping phenotypic and genotypic characters. Field biologists 

have found it increasingly difficult to place 

some individual animals into a species category 

because of the range of genetic and color variation observed in natural populations. As a 

result, sometimes an animal must be recorded to 

genus, species complex, or as “unknown” in 

field notes.

One of the best ways to identify salamander and frog larvae, in addition to color and 

morphology, is to examine their habitats and the 

times of year they are found. This can most easily be done through a comparative table. Morphological and life history characteristics are 

listed in tables 1 and 2 to help field biologists 

identify the species that are being examined. 

These data can be used in conjunction with the 

information in Dodd (2004).

SALAMANDERS

All salamanders in the Great Smoky 

Mountains have four limbs with four (Necturus, 

Hemidactylium) or five (all others) toes on each 

hind foot. They all have tails, lack dry scales 

covering the body (lizards have dry scales), and 

have skins that are moist or wet to the touch. 

The skins of a few species, such as Jordan=s 

Salamander (Plethodon jordani), are sticky 

because of glandular secretions, but only the 

Hellbender and Common Mudpuppy are truly 

slimy.

Biologists take two standard measurements with regard to length. The total length 

(TL) is the length of an animal from the tip of 

the snout to the tip of the tail. Because some 

salamanders lose their tails (or parts thereof) to 

predation, another common measurement 

recorded is the snout-vent length (SVL). SVL is 

measured from the tip of the snout to the posterior portion of the vent (the opening of the cloaca, the common receptacle for the digestive, 

excretory, and reproductive tracts). All scientific measurements are recorded in metric units, 

usually millimeters.

Salamander larvae sometimes are divided 

into two general groups, depending on morphology and the type of wetland in which they 

develop. The pond form (fig. 27A) is stout bodied, with long filamentous gills and a wide dorsal fin which extends well onto the body. Mole 

salamanders (Ambystoma), for example, have 

this type of larva. Pond larvae develop in still 

water, and use the extra surface area of the body 

and fin as aids in swimming. Stream larvae 

(Eurycea, Pseudotriton) are slimmer than pond 

larvae, with more streamlined bodies, shorter 
30

gills, and a narrower tail fin that does not extend 

onto the body (fig. 27B). These larvae usually 

live in swift flowing water, where extra surface 

area on the body would be a distinct disadvantage.

A number of useful characters are available which can be used to identify salamanders 

to genus or family. A few illustrative examples 

are provided, but more detailed comparisons are 

found in Dodd (2004) under the heading “Similar Species.”

Desmognathus: All dusky salamanders 

have a light line which extends from the back of 

the eye to the angle of the jaw. The duskies also 

have well-developed muscles on the sides of 

their heads. They need these muscles to raise 

the upper jaw in order to open their mouths, 

since the lower jaw is fused to the skull.

Gyrinophilus versus Pseudotriton: 

Although these colorful salamanders are superficially similar in appearance, Spring Salamanders 

(Gyrinophilus) have a canthus rostralis, a large 

white line bordered by black lines, that runs from 

in front of each eye to the nostrils. Salamanders 

of the genus Pseudotriton do not have this line. 

Spring Salamanders use the canthus rostralis as a 

“gunsight” to zero in on prey.

Plethodontidae versus all other salamander families: All lungless salamanders have 

a nasolabial groove that extends from each nostril to the upper jaw. The nasolabial groove 

transmits chemicals to the salamander from the 

substrate; no other salamander family has this 

groove.

FROGS

Like most salamanders, frogs have four 

legs with four toes on the front limbs and five 

toes on the rear limbs. The hind limbs are much 

larger than the front limbs, and are used to propel the body when walking, hopping, or jumping. Frogs are measured in TL, that is, from the 

tip of the snout to the end of the body between 

the hind limbs (that is, at the end of the 

antlerlike gills

costal grooves

tail fin

gill fimbriae dorsal fin

gill rachis

gular flap

(A) Pond form

(B) Stream form

Figure 27. Body morphology of a salamander larva: (A) pond form; (B) stream form.
31

urostyle). Of course, there are many other 

measurements which could be made, such 

as the length of the various sections of the 

hind limb, but these data generally are not 

important in amphibian monitoring-programs 

except in studies of fluctuating asymmetry 

(Alford and others, 1999, but see McCoy and 

Harris, 2003).

Tadpoles are morphologically complex. 

As with salamander larvae, there are two general tadpole body types, the pond type and the 

stream type. Pond-type tadpoles have deeper 

bodies and higher tail fins than do stream-type 

tadpoles. Structures important in the identification of tadpoles are labeled on figure 28. The 

Narial aperature

Spiracle Anus

MTH

(A) Lateral view

TMH

Oral disc

TL

BL TAL

IOD

IND

TMW

(B) Dorsal view

BL

IND

IOD

MTH

TAL

TL

TMH

TMW

body length

inter orbidal distance

maximum tail height

total length

tail muscle height

tail muscle width

inter narial distance

tail length

EXPLANATION

Figure 28. Body morphology of a tadpole.

oral disk consists of the mouth parts; the narial 

aperture is the opening to the nostrils; the 

spiracle is the opening from the gills (water is 

taken in through the mouth, passes over the 

gills, and is expelled via the spiracle); the anus 

is the opening from the digestive tract. The total 

length (TL) consists of the body length (BL) 

and tail length (TAL). Sometimes additional 

morphological measurements are taken, such as 

the maximum width of the tail musculature 

(TMH) or the maximum tail depth (MTH). The 

location and size of these characters, or their 

ratios in relation to one another, are useful in 

identifying what otherwise appears to be just 

another drab, olive-green, or black tadpole.
32

A-1

SM

MP

LJ

PL LP

A-2

Not

Emarginate

Labial teeth

Emarginate

AL

AL

A-1 and A-2

A-2 GAP

LJ

LP

M

MP

OD

PL

P-1, P-2, P-3

SM

UJ

A-2 Gap

UJ cuspate

P-1

P-3 P-2

anterior (upper) labium

first and second upper tooth rows

medial gap in second anterior tooth row

lower jaw sheath

lateral process of upper jaw sheath

mouth

marginal papilla

oral disk

posterior (lower) labium

first, second, and third posterior (lower) tooth rows

submarginal papilla

upper jaw sheath

EXPLANATION

Figure 29. Oral disc (mouthparts) of a tadpole.

The tadpoles of different 

species of frogs often appear identical to one another, but the structure of their mouthparts readily 

separate them. Biologists may 

need to examine mouthparts to 

determine which species is in 

hand. For this reason, a diagram 

has been included of tadpole 

mouthparts is provided in 

figure 29. The nomenclature follows Altig and McDiarmid (1999). 

The location, number, and degree 

of separation among labial teeth 

and papillae are important characters for identifying tadpoles. 

Examining tadpole oral disks 

(sometimes incorrectly termed 

“teeth”) also gives researchers an 

opportunity to check the health of 

the tadpole. For example, the 

horny jaw sheaths drop out when 

the tadpole is exposed to certain 

toxic compounds and to the dangerous disease, chytridiomycosis. 

However, tadpoles should not be 
33

held too long before examination or preservation, since some tadpoles may shed denticles in 

the laboratory.

As with salamanders, there are certain 

useful defining characteristics that help to identify certain superficially similar animals. Some 

of these are listed below (also see “Similar Species” in Dodd, 2004).

Bufonidae versus Pelobatidae: Frogs in 

both of these families are terrestrial. However, 

the true toads (Bufo) are dry-skinned and 

“warty,” and have prominent cranial crests and 

parotoid glands. The spadefoot toads (Scaphiopus) are smooth-skinned, lack cranial crests and 

parotoids, and have a sharp digging spade on 

their hind feet.

Hylidae versus other frog families: all 

hylid frogs (Acris, Pseudacris, Hyla) in the 

Great Smokies have slightly to completely 

expanded toepads, but only in the treefrogs 

(Hyla) are these greatly expanded for climbing; 

the other hylids are mostly ground-dwelling 

(however, note that Spring Peepers, Pseudacris 

crucifer, often call from the trees from late fall 

to early spring before descending to breeding 

ponds).

Rana palustris versus R. pipiens: these 

very similar frogs are both green and spotted. In 

R. palustris, the spots are squarish, paired and 

of nearly equal size, whereas in R. pipiens they 

are smaller, rounded, and more randomly scattered on the frog=s back.

Additional Information

Information on the etymology, identification of adults, larvae, and eggs, similar species 

and how to differentiate them, taxonomic problems, distribution both within the Park and elsewhere in North America, life history, abundance 

and status, and remarks on interesting aspects of 

the biology of the species are found in Dodd 

(2004). Data on 44 amphibians are presented, 

including information on species no longer 

thought present (for example, Aneides aeneus) 

or which were reported historically from the 

Park, but whose actual occurrence may be 

doubtful (Acris crepitans). Distribution maps, 

color photographs of amphibians from the Park, 

and original color illustrations accompany each 

account.
34

MONITORING PROGRAMS

Why Monitor Amphibians?

The problem of declining amphibian 

populations has been recognized worldwide, 

with credible reports of diminishment or disappearance of amphibians from many regions and 

habitat types. No single cause for declines has 

been demonstrated, although acid precipitation, 

environmental contaminants, introduction of 

exotic predators, disease agents, parasites, and 

effects of ultraviolet radiation have been suggested as factors in declining numbers. Indeed, 

no one cause may be implicated, and several 

factors may interact in such a manner as to 

threaten populations (Carey and Bryant, 1995). 

A major factor in the loss of amphibian populations has been and continues to be the loss of 

habitat. The severity and apparent complexity 

of the problem led the National Park Service in 

1997 to list amphibian declines as among its 

highest priority research and information needs.

In terms of its significance to amphibians, 

the Great Smoky Mountains National Park is 

more important than almost anywhere else in 

North America. Thirty-one species of salamanders have been recorded in the Park, and 

that number could conceivably increase as 

molecular genetic techniques are used to 

unravel the complex relations among 

populations. Of particular note are the salamanders of the family Plethodontidae, a largely 

North American group that has a center of evolution and distribution in the southern Appalachians (Dodd, 2004). Jordan=s Salamander 

(Plethodon jordani) is known to occur only in 

the Park, and the salamander fauna is believed 

to represent several evolutionary series progressing from the more aquatic species to those 

which are almost totally terrestrial. Thirteen 

species of frogs and toads are historically 

reported to inhabit the Park. The biological 

importance of the Park has been recognized in 

The problem of declining amphibian populations has been recognized worldwide....

its designation as an International Biosphere 

Reserve. Although no other region and no other 

National Park shares the wealth of amphibians 

found in the Great Smokies, the entire southern 

and midsection of the Appalachian chain is 

characterized by a high diversity of amphibians, 

and inventories and monitoring protocols developed in the Great Smokies may be applicable to 

National Park Service, U.S. Forest Service, 

Nature Conservancy, or other properties in the 

Appalachians.

Several known stressors potentially affect 

amphibians in the 2,071.2 km2

 (521,000 acre) 

Park (reviewed by Dodd, 2004). Air pollution, 

particularly long-distance pollution from cities 

in the nation's mid-region, is a nationally recognized problem. Reduced visibility, damage to 

plants, and fish kills are documented to be associated with sulfurous and nitrogenous compounds and atmospheric ozone. Low pH is 

known to have affected survivorship in at least 

one aquatic salamander species in the Park. 

Exotic pathogens and parasites have seriously 

affected some forest communities, with 

unknown effects on ecosystems. Finally, the 

pressure of ten million visitors per year--more 

than any other National Park--seems relatively 

benign, but could potentially have subtle effects 

on sensitive amphibian populations. The existence of these and other unknown stressors suggest that an inventory and a monitoring program 

are needed to ensure the protection of amphibian populations.

Amphibian Research and Monitoring 

Initiative (ARMI) — In 2000, the President of 

the United States and Congress directed Department of the Interior (DOI) agencies to develop a 

plan to monitor the trends in amphibian populations on DOI lands and to conduct research into 

possible causes of declines. The DOI has stewardship responsibilities over vast land holdings 

in the United States, much of which is occupied 

by or is potential habitat for amphibians. The 

U.S. Geological Survey (USGS) was given lead 

responsibility for planning and organizing this 

program, named the Amphibian Research and 

Monitoring Initiative (ARMI), in cooperation 
35

with the National Park Service, U.S. Fish and 

Wildlife Service, and Bureau of Land Management. Results of the monitoring program will be 

available to cooperators, land managers, the 

scientific community, and the general public. 

ARMI’s Internet site is:

http://edc2.usgs.gov/armi/

National Park Service (NPS) — Recent 

legislation (National Parks Omnibus Management Act of 1998) and policies of the National 

Park Service require that park managers know 

the condition of natural resources and that they 

monitor long-term trends in those resources. To 

comply with legal and policy requirements, the 

NPS Inventory and Monitoring Program 

focuses on attaining the following major 

long-term goals: (1) establish natural resource 

inventory and monitoring as a standard practice 

throughout the NPS that transcends traditional 

programs and activities; (2) inventory the natural resources and park ecosystems under NPS 

stewardship to determine their nature and 

status; (3) monitor park ecosystems to better 

understand their dynamic nature and condition 

and to provide reference points for comparisons 

with other, altered environments; (4) integrate 

natural resource inventory and monitoring 

information into NPS planning, management, 

and decision making; and, (5) share NPS 

accomplishments and information with other 

natural resource organizations and form partnerships for attaining common goals and objectives. Information on the National Park Service 

Inventory and Monitoring Program can be 

found at: http://www1.nature.nps.gov/im/monitor/index.htm #Legislation, and in publications 

by Silsbee and Peterson (1991) and Peterson 

and others (1995).

All Taxa Biodiversity Inventory 

(ATBI) — A research effort designed to compile a comprehensive inventory of all life forms 

in Great Smoky Mountains National Park, 

ATBI is sponsored by Discover Life in America, a private nongovernmental organization 

working in partnership with the NPS. The initiative has a goal of completing the inventory in as 

few as 10 years and is, therefore, an intensive 

undertaking. Before the project is completed, it 

will employ the expertise of taxonomists, data 

specialists, zoologists, botanists, and ecologists, 

among others. Once completed, the ATBI will 

provide baseline data from which to measure 

species change through time. ATBI=s objectives 

are to: (1) complete a comprehensive “checklist” of life forms in the Park; (2) gather data to 

create range maps for each Park species; 

(3) compile natural history information on each 

species, including its relative abundance, its 

response to various climatic conditions, photographs of each of its life stages, its role in the 

greater ecosystem, its relationship with other 

species, and digital recordings of its calls or 

sounds; and, (4) organize the information gathered and make it available to scientists, educators, land managers, students, and all other 

interested parties via the Internet and other 

media. More information can be found at: 

http://www.discoverlifeinamerica.org

Things to Consider During Planning

There are at least 10 major items which 

need to be addressed before starting an inventory or monitoring program for amphibians, 

especially when under financial or personnel 

constraints. These are discussed briefly below 

and, in some cases, more extensively elsewhere 

within the guide.

1. There are many amphibians in the southern 

Appalachians and the southeast. A total of 

31 species of salamanders and 13 species of 

frogs have been recorded historically as 

occurring in the Park. Extending the area of 

interest to the greater southern Appalachians, 

the figure increases substantially, by 21 salamanders and 1 frog, because of the high levels of endemism of many salamander 

species. Extending the area of interest even 

more, there are approximately 85 species of 

salamanders and 58 species of frogs within 

the southeastern United States (or 49.6 percent of the species in the entire United 

States). This figure does not include different 

subspecies, nor does it include the many 

genetic variants that have been described.

2. The systematic status of many species of 

southeastern amphibians is in a flux. It is 

likely that there are a number of new and 

unrecognized species of amphibians in the 
36

southern Appalachians, particularly among 

the salamanders. In addition, there is considerable debate among salamander taxonomists over what constitutes a species in 

terms of genetic uniqueness, phylogeny, 

and reproductive compatibility. Particularly 

in the genera Plethodon and Desmognathus, many new “genetic” species have 

been described in recent years, especially in 

the southern mountains. Unfortunately, 

morphology and coloration may be only of 

limited assistance in identification; many 

individuals are impossible to distinguish 

phenotypically in the field. There also are 

areas where considerable introgression or 

hybridization occurs, especially in the 

Great Smoky Mountains. This has led to the 

recognition of species complexes (for 

example, the slimy salamanders of the 

Plethodon glutinosus complex), or even of 

size-based guilds among the dusky salamanders (Desmognathus). As systematists 

closely examine other genera (Eurycea, 

Pseudotriton), the situation will probably 

become more complicated. Systematic certainty may be no better in the frog world, 

especially in the genera Pseudacris and 

Rana, although the taxonomy of frogs 

within the southern Appalachians will probably remain stable.

3. Species and life stages are sometimes difficult to distinguish. Even experienced herpetologists sometimes have difficulty 

identifying adult amphibians, and eggs and 

larvae pose special identification problems. 

Color and morphology vary considerably 

among individual amphibians. 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 naturalists who are 

knowledgeable concerning identification is 

diminishing. There are very few current 

color guides to amphibian eggs and larvae, 

even on a local basis.

4. Amphibians have complex life cycles. 

Because of the extremely varied life histories of many amphibians (see Life History), inventory and monitoring programs 

must consider such variation when planning 

when and where amphibians will be 

monitored, and what biases may be associated with interpreting sampling results.

For example, egg mass counts might tell 

a researcher about the number of egg 

masses deposited and, therefore, the number of females that reproduced that year. 

Egg mass counts cannot be used to determine population size (often used as a 

measure of status), however, unless the 

operational sex ratio (that is, the sex ratio of 

adults that actually bred successfully) is 

known for that year. This ratio is usually 

assumed to be 1:1, but if it is not, estimates 

of population size could be in error by 

several orders of magnitude. Also, not all 

individuals breed every year and, thus, population size at a breeding pond may not be 

indicative of overall population size. Even 

with such data available, population sizes 

still cannot be estimated inasmuch as the 

ratio of juveniles to adults is not known for 

most species. In addition, counting egg 

masses says nothing about whether reproduction was successful, since a variety of 

factors (disease, desiccation, predation) can 

interact to prevent hatching and metamorphosis. Consequently, it might be possible 

to count large numbers of egg masses, yet 

have none of them actually result in juvenile 

recruitment to the population. Status and 

long-range impacts to the population could 

be easily misinterpreted.

When inventorying and monitoring 

amphibians with complex life histories, 

multiple sampling techniques may be 

required, and status interpretation must be 

restricted to the sector of the population 

actually sampled. This rather obvious 

approach is often ignored, as authors often 

make general statements as to status and 

trends when only a portion of the animal=s 

life cycle was sampled.

5. In the field, detectability of amphibians is 

likely influenced by the following variables, 

to a greater or lesser extent, depending on 

species. Some of these variables include:

Annual cycles of reproduction–The 

reproductive season may be prolonged, or 
37

extend for only a few days or weeks. Some 

amphibians may be effectively sampled 

only during the breeding season 

(Ambystoma sp., Hemidactylium, many 

frogs), whereas breeding females of other 

species may disappear underground to 

brood eggs (Plethodon) and thus be undetectable. 

Seasonal events (cold, drought, heat, 

storms) that are usually unpredictable–

Cold, heat, and drought generally make 

amphibians more difficult to find, whereas 

tropical depressions and hurricanes, with 

their heavy rains, may actually bring 

amphibians to the surface in incredible 

numbers.

Diurnal versus nocturnal activity–

Many amphibians are more conspicuous at 

night, when they leave hiding places to forage, than they are in the day. This is true for 

both terrestrial and aquatic species.

Air, water, and substrate temperature–

Amphibians often have rather narrow tolerances or preferences for particular air, 

water, or substrate temperatures. Some species prefer rather cool temperatures (for 

example, salamanders living at high elevations, and the winter-breeding frogs), 

whereas others prefer the warm temperatures of summer. Since temperature 

changes with elevation (Dodd, 2004), activity patterns of broadly distributed species 

tend to change seasonally with an increase 

in elevation.

Soil moisture and rainfall–Terrestrial 

amphibians are active when soils are moist 

and during rainfall, much more so than 

when soils are dry. Breeding movements 

may be triggered by a combination of seasonal gonadal development, favorable temperature, and rainfall.

Relative humidity– High humidity 

favors amphibian activity; low humidity 

depresses activity.

Barometric pressure– Barometric pressure is indicative of changing weather conditions; a falling barometer is associated 

with weather fronts and rain, and a rising 

barometer is associated with clearing or fair 

weather. Therefore, a change in barometric 

pressure may influence amphibian activity 

patterns and, thus, detectability.

Cloud cover/moon brightness–

Amphibians tend to be more active on 

cloudy nights when humidity levels are 

higher than they are on clear nights. A 

bright moon tends to inhibit activity, since 

predators may be more effective at detecting prey on bright nights. 

Prey availability– Amphibians are likely 

to be more abundant in areas with a high 

diversity of prey items than in areas depauperate of prey. A few amphibians (Hellbenders) have specialized diet preferences. 

When prey are absent or scarce, specialist 

feeders will also be scarce despite the 

otherwise seemingly appropriateness of 

habitats.

Note that many of the variables discussed above change daily, seasonally, or 

annually (for example, during El Niño 

versus La Niña years).

6. Species and populations occur in a landscape. Some amphibian species are 

extremely localized geographically 

(Ambystoma opacum in the Great Smokies), whereas others are very widespread 

(Desmognathus quadramaculatus). Populations may be geographically isolated to an 

extreme degree (cave species or the 

crevice-dwelling Aneides aeneus), occur 

very patchily in a larger landscape, occur in 

a metapopulation structure (Bufo) with 

considerable (or little) interchange between 

or among metapopulations, or occur over 

hundreds of square kilometers of deciduous 

forest where it is difficult to define the 

limits of a population (many Plethodon). 

Individuals may be naturally rare or exceptionally abundant. Because a species is 

unusual or difficult to sample, is not a reason to bypass its study. Some of the most 

specialized amphibian species are those 

biologists know have declined or are imperiled in the southeastern states.

Although some populations may be huge 

(some terrestrial woodland salamanders, 

Plethodon, for example), others seem 

small, isolated, and vulnerable 

(crevice-dwelling, cave, or ravine species). 
38

Little is known about how and when these 

species disperse or about what mechanisms 

allow for the long-term persistence of small 

populations. Perhaps individuals move 

more than is recognized; even rare immigration is sufficient to ensure genetic 

exchange 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 suggests that generalizations about persistence will not be easily 

forthcoming.

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, disappear for 

years, then reappear. For example, populations of European Rana 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. Detectability may be influenced by 

weather (drought) even if populations are 

stable. Not much is known concerning the 

stability or fluctuation of semi-aquatic and 

most aquatic species and populations, especially in the southern Appalachians.

Still, biologists have enough 

data to advance hypotheses 

about the persistence and stability of amphibian populations, 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, whereas species 

...biologists have enough data to advance 

hypotheses about the persistence and 

stability of amphibian populations...

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 of 

species living in stable environments might 

be more cause for concern than declines in 

species living or breeding in fluctuating 

environments, unless the fluctuating environments are highly isolated. In this case, 

isolation may prevent recolonization from 

source populations and, thus, lead to 

declines throughout the landscape.

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 (as sources and 

sinks). Yet herpetologists understand little 

of these processes in southern Appalachian 

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, colonization and 

other forms of interpopulation movements 

may not move in a straight line overland. 

Animals might follow sinuous topography, 

watersheds, streams and rivers, or even subsurface passages.

Populations of amphibians certainly 

experience natural turnover (recruitment, 

mortality), but little is known about this 

process or how long it takes for any southern Appalachian species. Just because some 

individuals have the potential 

for considerable longevity 

does not mean that populations 

turn over slowly. Biologists need 

information on the generational 

times for various species.

9. Amphibian sampling techniques. There are as many ways to 

sample amphibians as there are 

amphibians (see Sampling Techniques). 
39

Each technique has its own underlying 

assumptions, biases, and limitations. Until 

relatively recently, these biases were unrecognized, not discussed, or simply ignored. 

Currently, sampling protocols have been 

receiving a great deal of experimental 

examination. 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.

10. The human-based constraints on sampling, 

inventorying, and monitoring amphibian 

populations on Federal lands must to be 

considered at the outset. These include: 

Money (equipment, personnel, emergencies, meetings, data analysis, publication) – The single biggest limitation 

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

• Sweep samples--for larvae

• Call surveys--

breeding or territorial adult frogs

• Egg mass counts

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 litterbags 

• Automatic audio data loggers--

for recording calling frogs

Intensive passive sampling

(labor, time, and financially expensive). 

• Traps and fences must be checked regularly, generally daily, for accurate 

results and to prevent mortality. 

• Traps (aquatic or terrestrial): funnels, 

bottles, minnow, wire basket

• Drift fences, with pitfalls and/or funnel 

traps, sometimes in conjunction with 

PVC pipes or coverboards

affecting inventory and monitoring projects 

is the amount of money available to conduct the programs, which ultimately will 

determine the number of researchers hired, 

the type of techniques used, the number of 

species monitored, and the number of locations visited. Inventory and monitoring programs should be designed to make the best 

use of the available funding to ensure scientific rigor, rather than try to be Aall things to 

all people.@

People (principal investigator, experienced field crews, biometricians, GIS, 

administrative support, field support) –

Highly qualified researchers and field technicians are absolutely essential for conducting inventory and monitoring programs. 

The identification of amphibians in the 

Great Smoky Mountains and elsewhere in 

the southern Appalachians is often difficult, 

and there is no substitute for experienced 

judgement. Resource managers should not 

assume that field assistants can be trained 

easily and quickly, or that volunteers can 

take the place of experienced biologists. 

Just as few persons would expect ecologists 

to conduct genetic analyses, current field 

research is a collaborative effort needing a 

variety of experts. When planning an inventory or monitoring program, agreements or 

arrangements need to be in place to ensure 

that field researchers have the needed biometric, landscape, and other types of support necessary for data analysis and 

interpretation.

Time – Inventory and monitoring programs take time to carry out. For amphibian 

monitoring programs, a minimum of 

10 years of data collection is not unreasonable to begin to understand population status and to measure the extent of variation 

associated with sampling data. Sampling 

time is dependent upon the life history 

characteristics of the species in question. 

Highly qualified researchers and field technicians are absolutely essential for conducting 

inventory and monitoring programs.
40

For example, a monitoring program might 

provide reliable trend-analysis data for a 

short-lived species if sampling was conducted every year for 10 years at locations 

throughout the species= range within the 

Park; for a long-lived species, the duration 

of sampling might have to extend for 20 to 

30 years before researchers could be confident in recognizing trends. In addition, 

trends resulting from human perturbation 

sometimes are difficult to separate from 

natural, often stochastic, population 

changes, except during catastrophic population collapse. It might be difficult to separate human-caused change from natural 

population variation without a long-term 

data set. Unfortunately, conflicts may arise 

when answers are needed by resource managers (for example, “We need to know the 

status of the Park=s amphibians for the 

annual report”). However, resource managers must recognize that short-term projects 

are ineffective and may give misleading 

results. Inventory and monitoring programs 

need time and patience. 

Safety – The minimum number of persons necessary to conduct amphibian field 

research involves two-person field crews. 

This is to ensure safety in case of injury, 

accident, or other medical emergency. 

Assume that emergencies will occur. Field 

crews should carry radios or cell phones 

and emergency first aid kits. Both heat 

stress and hypothermia are possible when 

sampling amphibians over long time periods in the southern Appalachians. Yellowjackets, venomous snakes, and bears are 

other park denizens requiring occasional 

attention.

Logistics – Can researchers get to locations with the people and equipment in a 

reasonable amount of time and effort? 

Given logistical constraints, how many 

sites can be sampled and over what area? 

The failure to consider logistical constraints is one of the most common errors 

when setting up inventory and monitoring 

programs. 

Regulations (permits, access, restrictions on research techniques, collecting) – 

Regulations can impede research results 

and limit the types of data collected. 

Researchers need to clearly understand the 

limitations imposed upon them by regulations, whether local, state, or national. Likewise, administrators need to recognize that 

some regulations can impede scientific 

progress. In some cases, it may be impossible to obtain scientific data given impositions upon research access or techniques.

Collaborations (intra-agency, Federal, 

state, other researchers, land managers) – 

Biologists working on amphibian inventory 

and monitoring programs should be knowledgeable about previous research and keep 

other researchers informed of their 

progress. When possible, ongoing research 

should be incorporated into the inventory or 

monitoring program to facilitate data sharing and partitioning of resources. Agency 

personnel need to facilitate research, especially for congressional or departmentally 

mandated programs.

Administrative Policy (hiring restrictions, equipment-ordering procedures, contracts) – Administrative delays need to be 

anticipated and alternative plans or policy 

established to allow science crews to be in 

the field conducting research when the 

animals are likely present.

Species and Locations to Monitor

Of the 44 amphibian species historically 

reported from the Park, two species (Green 

Salamander, Northern Cricket Frog) probably 

no longer occur within the Park; one species 

(Northern Leopard Frog) may not occur, and 

four species (Mole Salamander, Common Mudpuppy [perhaps], Mud Salamander, Eastern 

Spadefoot) are so rare that designing a meaningful species-based monitoring program for them 

is impossible. However, two of these species 

(the Mole Salamander and the Eastern Spadefoot) are known only from the same locality 

(Gum Swamp), which is also a major amphibian 

breeding site within the Park. Monitoring the 

amphibians at this site may result in occasional 

observations of these two restricted and rare 

species. Likewise, a monitoring program developed for the Hellbender might result in 
41

additional captures of the Common Mudpuppy, 

thus making it feasible to sample both species 

simultaneously.

The following suggestions are made to 

facilitate monitoring the amphibians of Great 

Smoky Mountains National Park. It is unlikely 

that all species within the Park can be monitored every sampling year, although careful 

planning may help to increase the number of 

species monitored through time.

1. Concentrate on certain species, especially 

those that may be in biological decline elsewhere within their range or are limited in 

distribution within the Park. Some of these 

species are:

• Large stream and river-dwelling species: 

Hellbender.

• Pond-breeding species: Spotted Salamander, Marbled Salamander, Eastern 

Red-spotted Newt, Four-toed Salamander, 

Northern Green Frog, Wood Frog.

• Stream-associated species (especially 

with conspicuous larvae): Black-bellied 

Salamander, Blue Ridge Two-lined Salamander, Black-chinned Red Salamander, 

Spring Salamander.

• (Primarily) Terrestrial salamanders: 

Jordan=s Salamander, Southern Graycheeked Salamander, Northern Slimy 

Salamander, Southern Red-backed Salamander, Southern Zigzag Salamander, 

Imitator Salamander, Pigmy Salamander.

2. Concentrate on areas of special species 

richness, such as the Cane Creek drainage, 

Cades Cove (especially Gum Swamp 

(fig. 18), Gourley Pond (fig. 20), Methodist Church Pond (fig. 22), Stupkas Sinkhole Pond (fig. 30), Big Spring Cove (the 

Finley-Cane sinkhole ponds), and the highelevation spruce-fir forest (fig. 3).

3. Concentrate on problem areas. The only 

currently recognized problem area for 

Figure 30. Biologist sampling with sweep net in Stupkas Sinkhole Pond in Cades Cove.
42

amphibians in Great Smoky Mountains 

National Park is Gourley Pond in Cades 

Cove. Amphibians breeding at this site have 

contracted iridovirus infections, and large 

numbers of larvae have died. Because of the 

disease threat (Chinchar, 2002), this location should be monitored every year 

throughout the breeding and metamorphic 

season, about mid-March to late July, 

depending on water levels.

4. Periodically check areas of known occurrence for certain species. There are a few 

areas within the Park where certain salamanders and frogs are known to occur with 

regularity; these locations can be visited 

periodically to determine continued presence and, possibly, relative abundance. 

The following are examples: Long-tailed 

Salamanders in Gregorys Cave and at other 

cave entrances; Cave Salamanders in 

Stupkas Cave; Southern Zigzag Salamanders in Whiteoak Sink and in the uvala 

surrounding Bull Cave; Seepage Salamanders along the road bordering Hazel 

Creek; American Bullfrog tadpoles in 

Abrams Creek at the Abrams Creek Ranger 

Station; Eastern Narrow-mouthed Toads at 

Shields Pond (fig. 31). If sampled during 

appropriate seasonal and weather conditions, these species should be found at the 

locations mentioned; if not, it could be an 

indication of concern. Unfortunately, it may 

be difficult to interpret such present/not 

observed data without information on the 

same species outside the Park.

5. If particularly cost-effective monitoring 

techniques are available for certain species, use them. For example, all breeding 

male frogs in the Park emit loud calls to 

attract females. Species that are extremely 

difficult to find at most times of the year, 

such as the Upland Chorus Frog (Pseudacris feriarum), can be readily detected 

calling on a wet spring night throughout 

Figure 31. Biologist looking for tadpoles at Shields Pond in Cades Cove.
43

Cades Cove. The presence and relative 

abundance of other breeding frogs that are 

spatially limited within the Park (such as 

the Eastern Narrow-mouthed Toad, Gastrophryne carolinensis, at Shields Pond in 

Cades Cove; fig. 31) can be detected by 

using automated call-monitoring devices 

without the continued presence of observers. As another example, the presence of 

certain salamander larvae can be detected 

passively using inconspicuously placed leaf 

litterbags. Larval Spring and Black-chinned 

Red Salamanders are detected in higher 

numbers using these bags compared to 

other search methods.

Choosing Sampling Sites

Pond-woodland pool breeding 

amphibians – If researchers decide to 

monitor the pond-woodland pool breeding 

amphibians within Great Smoky Mountains 

National Park, no great difficulty is encountered. This is because there are so few known 

locations that visiting each site two or more 

times per year can be planned very easily. One 

visit should be planned in the early spring (late 

March to mid-April), with a second visit in 

early summer (late May to mid-June). Coupled 

with at least one or two call surveys in Cades 

Cove and periodic call surveys at other locations, biologists should be able to determine 

whether most species are present, obtain counts 

of egg masses, and categorize the abundance of 

calling males. Because of the existing disease 

threat, Gourley Pond should be visited at least 

once every 3 to 4 weeks from February/March 

to July/August.

Large stream and river-dwelling 

amphibians – The Hellbender is the sole large 

stream- or river-dwelling species to be monitored in Great Smoky Mountains National Park. 

The largest population inhabits Little River 

from the Park entrance at Townsend for several 

kilometers within the Park, although the maximum distance upstream has not been determined. Smaller populations are found in lower 

Deep Creek and in the Oconaluftee River. The 

Little River population would, therefore, be the 

most important population to monitor annually. 

Periodic sampling should be conducted at the 

other locations and in potential habitat elsewhere within the Park (see Nickerson and others, 2002).

Streams and creek-dwelling 

amphibians – Depending on how precisely 

watersheds are defined, there are at least 

25 watersheds within Great Smoky Mountains 

National Park, totaling > 3,400 km of 

streambed. Nearly each meter of every stream 

likely contains salamanders. Sampling the 

amphibian fauna of these streams depends 

largely on: (1) objective (certain species or 

areas of interest); (2) money and personnel 

(how many field crews are available and can be 

hired); and, (3) time available to conduct the 

surveys. Obviously, it is necessary to define 

these limitations prior to undertaking a stream 

monitoring program. When deciding where to 

conduct a stream/creek amphibian monitoring 

program, researchers should decide first what 

they hope to accomplish. For example, if using 

“percentage of area occupied” (PAO) analyses 

(see Data Handling), many more sites can be 

sampled than by using intensive sampling or 

mark-recapture techniques. The objective will 

fit the analysis; this will be discussed in more 

detail in Data Handling.

Given the caveats of people and time 

constraints, it will be necessary to narrow the 

choice of stream locations to be sampled. Some 

ideas are listed in the following section. However, a biologist needs to remember that, as a 

rule, the more sites that are sampled, the greater 

confidence are the results. The goal of sampling 

is to determine reliable estimates of variance 

associated with capture or sighting probabilities, or with estimates of population size; variance estimates will be more reliable with a 

greater number of sites surveyed than with a 

small number of sites.

...there are at least 25 watersheds within 

Great Smoky Mountains National Park, totaling 

> 3,400 km of streambed. 
44

SAMPLING WATERSHEDS 

Limit sampling to a subset of watersheds: 

randomly pick watersheds to sample from 

throughout the Park. Each watershed is 

assigned a number and a computer program can 

then be used to select a random subset of the 

watersheds for survey. Streams to be sampled 

within the watershed are randomly selected in 

the same manner. The location of the exact part 

of the stream to be sampled can be specified 

randomly (very impractical in difficult-toaccess mountainous country) or stratified by 

stream order, elevation, vegetation type, access, 

or some other selective criterion. For example, 

biologists may limit their survey to second 

order streams between 900 and 1,400 m within 

1 mile by trail from a road access. A GIS can be 

used to generate the extent of such habitat with 

these criteria, locate potential sampling sites, 

and randomly select those to be sampled.

SAMPLING STREAMS

Limit sampling to a subset of streams: 

randomly select streams for sampling from 

throughout the Park. Each stream is assigned a 

number, and a computer program can be used to 

select a random subset of the streams for sampling. The location of the exact part of the 

stream can be specified randomly or stratified 

by stream order, elevation, vegetation type, 

access, or some other selective criterion, as in 

the example above. A GIS can be used to generate the extent of such habitat with these criteria, 

locate potential sampling sites, and randomly 

select those for sampling.

SAMPLING LOCATIONS

Specific locations can be selected for 

sampling, such as all streams draining into Tennessee, all streams draining into Cades Cove or 

Cataloochee Valley, or, all streams located on 

the western side of the Park. The same general 

procedure for site selection and stratification is 

followed. However, the more limited the area 

sampled, the more restricted generalizations 

about status must become. Researchers could 

not sample all the streams draining Mt. LeConte 

and then extrapolate their results concerning 

stream-dwelling salamander status to the entire 

Park, the eastern side of the Park, or even to 

nearby Mt. Guyot.

Terrestrial amphibians – Choosing 

terrestrial sites to sample for terrestrial salamanders is very similar to choosing stream sites, 

but without the streams. There is no welldefined physiographic feature, such as a watershed or stream course, with which to initially 

stratify the area to be sampled. Biologists are 

left with the questions: which species or 

amphibian community should be sampled, what 

habitats should be targeted, what areas should 

sampling be concentrated, and what degree of 

access is possible? Because the Park covers a 

large area (2,071.2 km2

), much of it in difficult 

terrain and without easy trail access, stratification of the terrestrial area to be sampled is absolutely necessary. How many sites can be 

sampled will depend on personnel, time available for sampling, and logistics. As with stream 

sampling, active sampling rather than passive 

sampling techniques will allow for more sites to 

be sampled, but the types of information that 

may be obtained will be correspondingly 

limited.

Unusual terrestrial amphibians – There 

are only a few salamanders that may qualify in 

this category, such as the Southern Zigzag Salamander currently known from only two areas 

within the Park (Whiteoak Sink; entrance to 

Bull Cave), and the cave entrance-inhabiting 

salamanders of Gregorys Cave, the Calf Caves, 

and Stupkas Cave (especially Long-tailed and 

Cave Salamanders). As with sampling pondbreeding amphibians, these sites could be 

checked annually to verify the presence of these 

species. Detailed studies, using mark-recapture 

techniques, would be necessary to establish 

population size and trends through time.
45

SAMPLING TECHNIQUES AND PROTOCOLS

In the section that follows, brief examples are listed of how certain techniques have 

been used to sample amphibians. As stated in 

Things to Consider During Planning, there 

may be vastly different amounts of time associated with using the different techniques, different reasons for choosing them, and different 

biases when interpreting the results. In every 

instance, researchers should quantify the 

amount of search time or sampling effort 

involved in the survey.

Active Sampling 

Time constrained – In this technique, a 

predetermined amount of time is set for sampling the area or habitat. The presence of different species and the number of individuals (or 

even sex and life stage—males, females, juveniles) observed are recorded. Visual encounter 

protocols are followed; that is, animals are 

counted as they walk over the forest floor or 

stream bottom, hide in crevices or cling to cave 

Figure 32. Turning logs in time constrained survey at Beech Flats.

walls, found by turning over surface debris 

(figs. 32, 33), heard calling, or captured in random dip (fig. 34) or sweep nets (fig. 30). The 

number of observers x total amount of time 

sampled is recorded. In terrestrial and aquatic 

situations, times may be set for 15 or 30 minutes, occasionally longer, depending on the 

number of observers and the amount or quality 

of habitat to be surveyed.

Example. A sampling protocol is set 

whereby three researchers hike along Noland 

Divide Trail for 30 minutes, conduct a 

30 minute time-constrained survey, hike 

another 30 minutes followed by another 

30 minute sample, and so on throughout the 

day. Four to six sites per day can be sampled 

with this method, depending on trail conditions 

and terrain. The sampling effort would be 

3 x 30 = 90 person-minutes at each site sampled. Sample data might be 3 adult D. imitator, 

5 P. jordani (2 males, 3 juveniles), and 1 subadult E. wilderae at site 1, with similar data 

recorded at every sampling location.
46

Figure 34. Dip netting for salamander larvae in 

Abrams Creek.

Figure 33. Terrestrial timeconstrained survey in thickly 

vegetated habitat at Balsam 

Mountain.
47

 What this tells the observer. Timeconstrained surveys provide information on: 

(1) species presence (but not absence) at the 

time of sampling; (2) life history information, 

such as when eggs are deposited, larval presence, and activity patterns; and (3) habitat information. Sampling effort is easily quantified.

Limitations. Detectability is influenced 

by all the factors listed in Things to Consider 

During Planning. Even if every attempt is 

made to standardize sampling (for example, by 

sampling at the same time of day and during the 

same time of year), environmental factors likely 

will be different and thus influence whether a 

species will be observed. Because environmental variables influence the number of animals 

observed, differences in counts over time may 

be more reflective of differences in environmental conditions during the sampling periods 

among years than changes in status. It is very 

difficult to determine any kind of trend based on 

periodic counts because it is unknown what the 

relationship is between the counts and actual 

abundance. In addition, there may be considerable variation in the ability of the field observers 

to locate and count animals; some observers may 

find animals easily, whereas others might have 

great difficulty finding amphibians. Observer 

bias, thus, could skew count data in a manner 

which has nothing to do with the actual abundance of the animals counted.

Area constrained – In this technique, a 

defined amount of habitat is selected for sampling. For example, researchers might choose to 

sample large, randomly selected plots (such as 

30 x 40 m plots; fig. 35); they might survey 

smaller plots (for example, 10 x 10 m plots) during a hiking survey; or they might survey a 

pond, wetland, or cave entrance, regardless of 

how much time is required. Plots may be singular or in groups (fig. 36). As above, the presence 

of different species and the number of individuals (or even sex and life stageBmales, females, 

juveniles) observed are recorded. Visual 

encounter protocols also are followed; that is, 

animals are counted as they walk over the forest 

floor or stream bottom, hide in crevices or cling 

to cave walls, found by turning over surface 

debris, heard calling, or captured in random dip 

or sweep nets. The number of observers x total 

amount of time sampled is recorded.

Example. Two persons search Gourley 

Pond for 67 minutes. The sampling effort is 

2 x 67 = 134 person-minutes. Sample data 

might be: larval A. opacum (> 50 observed), 

14 egg masses of A. maculatum, larval R. sylvatica (hundreds of tadpoles), 4 P. crucifer

heard calling.

 What this tells the observer. Areaconstrained surveys provide information on: 

(1) species presence (but not absence) at the 

time of sampling; (2) life history information, 

such as when eggs are deposited, larval presence, and activity patterns; (3) habitat information; and (4) in some cases, a very crude 

estimate of density (the amount of area ) number of animals). Sampling effort is easily 

quantified.

Limitations. Detectability is influenced 

by all the factors listed in Things to Consider 

During Planning. Even if every attempt again 

is made to standardize sampling, environmental 

factors likely will be different and thus influence whether a species is observed. Since environmental variables influence the number of 

animals observed, differences in counts over 

time may be more reflective of differences in 

environmental conditions during the sampling 

periods among years rather than changes in 

amphibian population status. As with timeconstrained sampling, it is very difficult to 

determine any kind of trend based on periodic 

counts because the relationship between counts 

and actual abundance is unknown.

Transects – Transect sampling can be 

conducted using simple visual encounter survey 

techniques, such as by walking a preselected 

line transect at night and counting all the salamanders seen, or it can be used in conjunction 

with passive sampling techniques, such as the 

placement of coverboards along a preselected 

survey line. When using transects, sampling 

locations are determined through a stratified 

random process. A survey line of a prescribed 

length is selected, and observers use the line as 

a base from which to make observations.
48

flowing stream

30 x 40 m Study Plot

Data

Logger

DISTANCE, IN METERS

0 5 10 20 30 40 15 25 35

0

5

10

15

20

25

30

DISTANCE, IN METERS

Blue flags located

at every perimeter

intersection

Pink flags located

at every interior

intersection

A surveyed in June,

B surveyed in July,

C surveyed in August

3, lots 30 x 40 meter p

surveyed once per year

Three study plots at one site

A

B

C

Data

Logger

Stream

Figure 35. Schematic of a 30 x 

40-meter sampling plot. The grid 

is marked off in 5-meter intervals. 

The outside of the grid is marked 

with blue survey flags, whereas 

the rows are marked with pink 

survey flags. A stream is included 

on the left margin of the plot, so 

that both stream and terrestrial 

salamanders may be surveyed. 

Automated data loggers (red dot, 

DL) can be installed to record air 

and water temperature and 

relative humidity. Researchers 

walk up the survey lines turning 

coarse woody debris, rocks, and 

leaf litter. In addition to information 

on the species, size, and age 

class of salamanders observed or 

captured, the distance from water 

also can be recorded. This gives 

an idea of the spatial distribution 

of species across the plot.

Figure 36. Diagram of the relationship of three 30 x 40-meter 

fixed sampling plots at a location. Plots need not be isolated. 

In this schematic, three plots are located along the course of a 

stream. Each plot is surveyed once per year during the 

summer, all in the same order (A in June; B in July; C in 

August), for the length of the study. A single data logger 

station is located at one of the plots.
49

Example 1. Researchers select 50 locations in the fir-spruce forest where transects of 

100 m length will be established. During the 

day, a starting point for the transect is selected. 

The direction of the transect is then determined 

from a set of random numbers from 1 to 360 

(based on the number of degrees in a circle). 

Using a compass and a 100-m survey tape, fluorescent tape is used to designate the survey 

line. After dark, two researchers walk along the 

transect line, 5 minutes apart, and count all the 

salamanders, categorized by species, observed 

in their flashlight beams. The distance from the 

starting point where the salamanders were 

observed also is recorded. Using two researchers allows for a measure of potential observer 

bias.

Example 2. A three-party survey crew 

samples the Little River for Hellbenders. The 

total amount of the river to be sampled is 

marked off in 100-m sections on a map, and ten 

100-m sections are selected for sampling based 

on a random numbers chart. At the river, a 

Figure 37. Stream sampling at Balsam Mountain.

starting point and an end point are marked using 

red survey flagging. Wearing wet suits, two 

observers snorkel along parallel transects about 

4 m from the shore and look for Hellbenders 

under rocks, ledges, and other underwater hiding places. Observations are relayed to the third 

researcher walking parallel to the shore.

Example 3. Researchers select 50 stream 

locations on the northern side of the Park for 

sampling; the locations are selected based on 

elevation and accessibility. At each location, the 

stream is marked off in 5-m transects for a total 

of 100 m of stream length. Using a random 

numbers chart, seven transects are selected for 

sampling. A two-person team turns over all the 

rocks and searches hiding places, beginning 

downstream and working upstream, capturing 

and measuring salamanders (fig. 37). They call 

out the data (species, sex, length, age-class) to a 

third researcher walking parallel to the stream 

who records the information (fig. 38).
50

Figure 38. Checking identification and recording data during stream sampling at Balsam Mountain.

Example 4. Researchers select 50 locations in the fir-spruce forest where transects of 

100 m in length will be established. A starting 

point for the transect is selected. The direction 

of the transect is then determined from a set of 

random numbers (from 1 to 360, based on the 

number of degrees in a circle). Using a compass 

and a 100-m survey tape, fluorescent tape is 

used to mark the survey line. At every 10-m 

increment, a series of eight coverboards are laid 

out in a grid parallel to the transect line (fig. 39). 

The coverboards are then monitored periodically for salamander presence (see Coverboards).

 What this tells the observer. Areaconstrained surveys provide information on: 

(1) species presence (but not absence) at the 

time of sampling; (2) life history information, 

such as when eggs are deposited, larval presence, size-class structure, and activity patterns; 

(3) habitat information; and (4) in some cases, a 

very crude estimate of density (for example, a 

minimum number of salamanders inhabiting the 

selected length of the stream surveyed). 

Sampling effort is easily quantified.

Limitations. Detectability is influenced by all 

the factors listed in Things to Consider During 

Planning. Even if every attempt is made to standardize sampling (for example, by sampling at 

the same time of day and during the same time 

of year), environmental factors likely will be 

different and thus influence whether a species is 

observed. Because environmental variables 

influence the number of animals observed, differences in counts over time may be more 

reflective of differences in environmental conditions during the sampling periods among years 

than changes in amphibian population status. It 

is very difficult to determine any kind of trend 

based on periodic counts, because it is unknown 

what the relationship is between the counts and 

actual abundance. On the other hand, the lifehistory information obtained using transect 

surveys may be valuable for understanding the 

basic biology and demography of the species 

sampled.
51

Sweep samples – Sweeping a large, 

small-mesh dip net through the water column or 

in submerged leaf litter in ponds or larger wetlands allows observers to capture amphibian larvae and sometimes breeding adults. Sample 

locations may be completely randomized or 

some measure of design can be incorporated 

into sampling, such as by sampling areas along 

pond margins every 10 or 15 m, depending on 

the circumference of the area to be sampled. 

Species richness, the number of larvae in each 

sweep, and the total number of sweeps are 

recorded.

Example. Two persons search the entire 

circumference around Gourley Pond by sweeping a dip net five times every 15 m. If the pond 

margin is 600 m, then 40 locations could be 

sampled and 200 sweeps could be made. The 

sampling effort is 200 sweeps. Sample data 

might be: 240 larval A. opacum; 6 egg masses of 

A. maculatum; and, 1,246 larval R. sylvatica. 

The amount of area sampled in relation to available habitat could be estimated visually.

 What this tells the observer. Sweep surveys provide information on: (1) larval species 

presence at the time of sampling; (2) life history 

information, such as when eggs are deposited 

and tadpole developmental stage; (3) habitat 

information, such as microhabitat preferences 

and distribution of various larvae; and, (4) in 

some cases, an estimate of density (number of 

Transect and Coverboard Survey

Environmental/physical data recorded here

0 50

48 coverboards, approximately 12 x 24 inches, placed perpendicular to a 50-meter transect line at 10-meter intervals.

Coverboards extend approximately 5 meters on either side of transect.

Figure 39. Schematic of a combined transect/coverboard survey design. A series of eight 

coverboards are located at the origin and thereafter at 10-meter intervals along a 50-meter transect. In 

this design, the boards are placed perpendicular to the transect. This survey design could be 

combined with a night survey, whereby a team of observers walks along the transect, spotlighting and 

counting salamanders. The boards would not be disturbed during such survey. Using dual observers 

at close intervals helps quantify observer bias. See text for layout details.

animals ) the amount of area sampled in reference to available habitat). Sampling effort is 

easily quantified.

Limitations. Detectability may be influenced by many of the factors listed in Things to 

Consider During Planning. Even if every 

attempt is made to standardize sampling, environmental factors (for example, water availability and depth; water temperature) likely will be 

different among sampling occasions and thus 

influence whether a species is observed. Since 

environmental variables influence the number 

of animals observed, differences in counts over 

time may be only reflective of differences in 

environmental conditions during sampling periods. As with time-constrained sampling, it is 

very difficult to determine any kind of trend 

based on periodic counts, because the relationship between counts and actual abundance is 

unknown. Also, the number of larvae observed 

may not reflect the number of breeding adults, 

or tell anything about future reproductive success and the rate of successful metamorphosis. 

....differences in counts over time may be 

only reflective of differences in environmental conditions during sampling periods.
52

For example, the wetland could dry 10 days 

after a sampling visit, and all larvae could 

perish.

Call surveys – All species of male frogs 

in Great Smoky Mountains National Park call to 

establish breeding territories and attract 

females. Species that may be quite difficult to 

find throughout most of the year can be readily 

heard at this time, their breeding sites identified, 

and relative abundances of adult calling males 

estimated. Call surveys are easy to conduct. A 

biologist simply periodically visits wetlands or 

drives park roads at night during the breeding 

season and records the locations of species 

heard calling. In very large choruses, it may be 

necessary to record abundance in terms of categories: 1 = 0 frogs calling; 2 = 1 individual calling; 3 = < 5 individuals calling; 4 = > 5 to 10 

individuals calling; 5 = > 10 individuals calling.

Areas appropriate for call surveys within 

the Great Smokies include the Cades Cove 

Loop Road and associated roads in Cades Cove, 

the road through Cataloochee Valley, Laurel 

Creek Road, Little River Road, lowland areas of 

Newfound Gap Road at Sugarlands and Smokemont, Big Cove Road, and the entry roads to 

Greenbrier, Cosby, and Deep Creek. Two methods may be used: (1) drive slowly and listen for 

frog choruses, or (2) conduct systematic 

searches using periodic stops with defined 

amounts of time for listening.

Example. Starting at the entry gate to 

Cades Cove Loop Road, drive slowly and stop 

every 0.5 miles. At each stop, turn off the 

engine, and listen for 5 minutes. Record the 

species heard and the compass direction from 

which the call is heard; possible breeding sites 

can be identified during daylight hours as time 

permits.

 What this tells the observer. Call surveys 

provide information on: (1) adult male presence 

at the time of sampling; (2) the dates and 

Call surveys must be conducted at multiple occasions during the potential breeding season.

environmental conditions when males call; (3) 

the location of breeding sites; and (4) an estimate of breeding male relative abundance can 

be attained through the use of the abundance 

categories. Sampling effort is easily quantified.

Limitations. Detectability may be influenced by many of the factors listed in Things to 

Consider During Planning. Even if every 

attempt is made to standardize sampling, environmental factors (for example, weather, temperature, rainfall patterns) likely will be 

different among sampling occasions and thus 

influence whether a species is heard. Since environmental variables influence the number of 

animals calling, differences among abundance 

categories over time may be only reflective of 

differences in environmental conditions during 

sampling periods. Thus, call surveys must be 

conducted at multiple occasions during the 

potential breeding season. Further, call surveys 

tell nothing about the presence and number of 

females and nonbreeding males, or whether 

reproduction was successful. Call surveys are 

best implemented where researchers have 

access by road; isolated breeding sites could be 

overlooked, or ignored when access is difficult 

(such as along lower Hazel and Eagle Creeks). 

Since frogs often call diurnally or during different intervals of the night (several hours after 

dusk or before dawn), species could be missed 

or relative abundances underestimated. One way 

to circumvent this problem is to use automated 

data loggers to periodically sample frog calls 

throughout the day and night.

Egg mass or nest counts – A number of 

amphibians (Spotted Salamander, Wood Frog) 

deposit globular egg masses that are readily 

identified and can be counted. Other species 

(Marbled Salamander, Four-toed Salamander) 

deposit eggs in terrestrial habitat on dry pond 

bottoms or in the vegetation bordering ponds. 

As the pond fills, the eggs are inundated and 

hatching occurs (Marbled Salamander) or the 

eggs hatch and larvae wiggle through the vegetation to reach the pond (Four-toed Salamander). Counting egg masses or nests should 

give an indication of reproduction during the 

sampling period. This method has been used in 

the Great Smokies by James Petranka and 

Charles Smith; Crouch and Paton (2000) have 
53

suggested that the method is an effective way to 

gage trends in Wood Frog population size and 

reproduction.

Example 1. Researchers visit Gum 

Swamp shortly after Wood Frogs have bred. 

Each separate egg mass can be identified and a 

flag placed next to it. Flags mark the distribution of the egg masses, are easily counted, can 

be left in place to follow reproductive parameters (for example, whether successful hatching 

takes place), and help to reduce observer bias 

(single observers can miss 10 percent or more of 

the egg masses (Crouch and Paton, 2000)). 

Because each female deposits one mass, the 

number of breeding females at a pond can be 

monitored through time.

Example 2. The dry pond basin at Gum 

Swamp can be searched in October when 

female A. opacum have deposited their eggs and 

are sitting over them until the autumn rains 

arrive. By carefully turning logs, researchers 

can locate nests, place flags in the ground adjacent to them, and obtain an idea of the number 

of nests and their spatial distribution. Numbers 

of females and males can be counted (see Dodd, 

2004, for sex determination criteria).

 What this tells the observer. Egg mass or 

nest surveys provide information on: (1) the 

number of females breeding successfully in a 

year; (2) the dates and environmental conditions 

when eggs are deposited; and (3) egg masses 

that can be followed through time to obtain an 

idea of the extent of successful reproduction. 

Crude estimates of the number of metamorphs 

produced can be obtained (number of egg 

masses x the percentage of masses with successful hatching x the mean number of eggs per 

mass). In the case of nests, the reproductive 

potential (number of nests x the mean number 

of eggs per nest) can be determined. Sampling 

effort is easily quantified as the amount of time 

spent searching an area.

Limitations. Counting egg masses 

assumes that there is one female per egg mass. 

This assumption seems to hold true for those 

species depositing large, globular, jelly masses. 

However, this assumption will not be valid for 

all species depositing eggs in nests (for example, the Four-toed Salamander) because nests 

may include the eggs of more than one female. 

Be sure to check information on life history 

(Dodd, 2004). Counting egg masses generally 

does not give an indication of the number of 

males or nonbreeding females (but see Crouch 

and Paton, 2000). Unless the hatching success 

of egg masses is recorded, counting egg masses 

will not provide an estimate of the number of 

metamorphs produced during the breeding 

season. Care must be taken not to disturb 

brooding females because nest abandonment 

virtually ensures reproductive failure. Although 

some species are more tolerant of disturbance 

than others, a nest should not be disturbed 

repeatedly.

Easy Passive Sampling

Coverboards – Herpetologists have a 

long history of turning over surface cover 

objects to look for terrestrial salamanders and 

reptiles. Coverboards are simply an extension of 

this search technique, albeit with a more formalized sampling design. Coverboards may be 

made of many types of materials (for example, 

wood, tarpaper shingles, plastic sheets), but the 

most common material is nonchemically treated 

plywood. The boards are cut into small sizes 

(for example, 20 x 25 cm; 35 x 35 cm; fig. 40) 

and placed in a grid of various design. Boards 

should not be too large, because the leaf litter 

Figure 40. Coverboards.
54

underneath them becomes dry in the center and 

discourages salamander residency. Pressuretreated boards should never be used.

In the Great Smoky Mountains, National 

Park Service personnel have used four boards 

placed within a few centimeters of one another 

at each sampling site along a long transect. 

Sampling sites might be located at 10-m intervals along the transect, such that a 50-m transect 

would have 24 coverboards placed along it (stations 0-5 x 4 boards/station). Coverboards must

be placed in location for at least a month prior 

to beginning a survey to ensure they age properly and provide secure hiding places. Ideally, 

coverboards should be set out in the autumn of 

the preceding year prior to sampling. Some 

researchers scrape the ground underneath 

coverboards to ensure that the area underneath 

is not too large to discourage residence or will 

not increase air flow. Coverboards should be 

checked once every week or two; too much disturbance will inhibit salamander occupancy.

Example. In a study of sampling techniques on the north side of Mt. LeConte, Hyde 

and Simons (2001) used two sizes of coverboards (three 13 x 26 cm; two 26 x 26 cm) 

placed at 10-m intervals along a 50-m transect 

(5 boards x 5 sampling stations = 25 boards/

transect). Using a stratified sampling design to 

locate transect sites, they sampled 101 locations 

and captured 1,224 salamanders over a 2-year 

period. Coverboards were only checked three 

times the first year, and four times the second 

year.

 What this tells the observer. Coverboard 

surveys provide information on: (1) species 

presence at the time of sampling; (2) life history 

information, such as data on size-class structure, reproduction, and activity patterns; and 

(3) habitat information. If used in conjunction 

with mark-recapture techniques, they also 

might be used to examine site fidelity, movement, and population size. Sampling effort is 

easily quantified (number of coverboards x 

number of days sampled).

Limitations. Capture probability is 

influenced by all the factors listed in Things to 

Consider During Planning. Even if every 

attempt is made to standardize sampling, 

environmental factors likely will be different 

and thus influence whether a species is 

observed. Because environmental variables 

influence the number of animals observed, differences in counts over time may be more 

reflective of differences in environmental 

conditions during the sampling periods among 

years than changes in status. It is very difficult 

to determine any kind of trend based on periodic 

counts, because it is unknown what the relationship is between the counts and actual abundance. Hyde and Simons (2001) found that 

counts of terrestrial salamanders in the Great 

Smokies were highly variable and that sampling 

variability and detectability were not constant 

among species or even habitat type. Recapture 

rates of marked salamanders also are notoriously low, making estimates of population size 

unreliable. Finally, coverboards may provide 

artificially favorable cover, although preliminary evidence suggests this capture bias may 

not be as serious as previously believed. Some 

size classes of terrestrial salamanders are more 

likely to use coverboards than other sizes (for 

example, data from Virginia suggest that 

hatchlings and juveniles are found less often 

under coverboards than they are under natural 

cover objects). Coverboards are labor intensive 

to cut and haul to a sampling site. They are 

subject to vandalism, and bears and pigs will 

readily turn them over or move them around.

PVC pipes – A method that has proved 

successful in the southeastern United States for 

monitoring treefrog (Hyla) populations is to 

place polyvinyl chloride (PVC) pipes in the 

ground or to mount them on trees (Boughton 

and others, 2000; http://www.fcsc.usgs.gov/

posters/Herpetology/Artificial_ Refugia/

artificial_refugia.html). The pipes are readily 

colonized by treefrogs, even during the nonbreeding season when the treefrogs are dispersed away from ponds. The placement of the 

pipes and their characteristics (diameter, structure, possibly color) are important. Frogs are 

captured most often in pipes of 3.8 to 5.0 cm 

(1.75-2 inch) in diameter located 2- to 4-m high, 

on a large trunked, deciduous, hardwood tree; 

they are captured much less frequently in pipes 

on tree trunks near the ground, in pipes of larger 

diameter, or in pipes located on pine trees 
55

(fig. 41). Pipes capped on the bottom to allow 

some standing water within the shaft and 

presumably to increase humidity also capture 

more frogs than pipes that are open on both 

ends. Free-standing pipes (91.4 cm; 36 inches) 

sunk directly in the ground near breeding ponds 

also are used by treefrogs. 

Example. A series of PVC pipes are to be 

placed around Gourley Pond to monitor the 

population of Cope=s Gray Treefrog (Hyla chrysoscelis). Twenty transects are established 

evenly spaced around the pond perimeter at its 

edge (fig. 42). Each transect consists of five 

pairs of pipes (N = 10/transect; total N = 200 

pipes) spaced 10 m apart, and radiates outward 

perpendicular to the pond=s edge, similar to the 

spokes of a wheel. The first two pairs are inground pipes, whereas the last three pairs are 

nailed to hardwood trees (if possible) at a 2-m 

height. Each pair of pipes consists of one 3.8- 

and one 5.0-cm pipe. The pipes on trees are fitted with bottom caps, with a hole drilled 9 cm 

above the base to allow drainage. Pipes are 

painted camouflage green on the outside for 

concealment, and each pipe is marked with a 

distinct number. Pipes are checked once a week 

from March through September. The number of 

frogs observed is recorded. Frogs could be 

Figure 41. PVC 

pipes on trees in 

Okefenokee 

National Wildlife 

Refuge.

marked via individual or cohort toe clips, or digitally photographed for identification. Recording the data separately for unmarked animals 

and recaptures is important, because results 

from other studies show that frogs take up residency within pipes.

 What this tells the observer. PVC pipe 

surveys provide information on: (1) species 

presence at the time of sampling; (2) life history 

information, such as when animals arrive at 

breeding ponds, how long they stay, sex ratios, 

size-class structure; (3) movement patterns 

while at the ponds; and (4) information on the 

direction and distance of dispersal. Sampling 

effort is easily quantified (number of pipes x the 

number of 24-hour periods sampled).

Limitations. The only species that can 

be monitored in the Park using PVC pipes is 

Cope=s Gray Treefrog. Even then, sampling 

results for this species have revealed mixed 

results at other locations where pipes have been 

used. In some areas, Cope=s Gray Treefrogs will 

use pipes as retreats, whereas in other areas they 

seem to avoid PVC pipes. Whether they will use 

PVC pipes in the Great Smokies is unknown. If 

simple presence data are needed, call surveys 

would be more appropriate, although PVC 
56

Pipes

POND

Tree

EXPLANATION

Typical transect with 5

stations, each with a pair

of pipes

Pond basin margin

Transect with 5 stations,

each with a pair of pipes

Station with 2 pipes in the

ground

Station with 2 pipes on a

tree

Figure 42. Schematic of a survey design using paired PVC pipes located at 10-meter intervals around a 

pond’s perimeter. The first set of pipes is located at the pond’s margin, and thereafter at 5- or 10-meter 

intervals perpendicular to the pond. The second set of pipes is located at the margin of the pond basin 

(dashed line). The first two sets of pipes are ground pipes (black dots), whereas the last three (gold dots) are 

located (preferably) on large-diameter deciduous hardwoods. Pipes are placed at a height of 2 meters on 

opposite sides of the trunk (red dots).

sampling might prove valuable if more detailed 

life-history information is required. PVC pipes 

are likely to be stolen or vandalized. Bears, in 

particular, seem to be attracted to PVC and will 

often bite it or carry pieces around.

Larval litterbags – One relatively new 

method for inventorying and sampling most 

stream-dwelling salamanders, especially larvae, 

involves the use of artificial refugia (leaf litterbags) placed in shallow streams (fig. 43). In 

2000, Waldron and others (2003) tested the 

utility of using litterbags to sample salamanders 

in Great Smoky Mountains National Park. 

Three transects of six litterbags each (two large, 

two medium, and two small) were placed in five 

small, medium, and large streams. A total of 

690 larval, juvenile, and adult stream-dwelling 

salamanders from 11 species were captured 

from June to November in the 90 litterbags. 

Sampling salamanders in small streams was 

most productive using large and medium-sized 

litterbags, although all bag sizes worked equally 

well in medium and large streams. The number 

of salamanders captured varied seasonally, with 

most captures occurring in June and July. The 

depth of bag submergence significantly influenced litterbag use by adult and larval salamanders, but had no effect on use by juvenile 

salamanders. The ease of deployment and nondestructive sampling methodology suggest that 
57

litterbags could be useful in determining salamander presence during large-scale inventory 

programs, especially when the time available 

for sampling a large number of individual sites 

is limited and when sampling for secretive or 

uncommon larvae, such as Pseudotriton or 

Gyrinophilus.

Example. Litterbags of two sizes (70 x 70 

and 90 x 90 cm) are constructed as outlined in 

Waldron and others (2003). In the field, three or 

four small rocks are placed in the netting to give 

the bag weight, then covered with leaves. Once 

filled with leaf litter, the corners of the netting 

are pulled together and tied with plastic cable 

ties to form a bag. Blue flagging is tied to the 

top of each bag so that researchers can easily 

locate bags in the field. Precautions are taken to 

prevent the loss of bags from fast-flowing water 

and flooding by placing one or two large rocks 

against or just downstream from each bag, and 

by tethering each bag to the nearest root, log, or 

large rock using monofilament fishing line.

Streams are selected using a stratified 

sampling protocol for size, location, and ease of 

Figure 43. Leaf litterbag in Little River.

access (see Sampling Streams). All streams are 

< 50 cm in depth at the sampling site. Sampling 

sites are spaced so that a watershed can be 

sampled in 1 day, allowing all of the sites to be 

completely sampled in 1 week. One 50-m 

transect is set up in each stream study area. 

Eight bags, four of each size class, are placed 

10 m apart along transects. The order of presentation of medium and large bags from 0 to 50-m 

is randomized along the transect. Litterbags are 

sampled biweekly from April through September. Prior to sampling each litterbag, the percentage of litterbag submergence under water is 

recorded. Bags are removed quickly from the 

stream and gently shaken over a white dishpan 

for approximately 15 seconds to remove salamanders (fig. 44). Adult, juvenile, and larval 

salamanders that fall into the dishpan are identified to species, measured for total length (TL, 

tip of snout to end of the tail) and snoutto-vent-length (SVL, tip of snout to the posterior end of the cloacal opening), and released. If 

field identification is not possible, individuals 
58

Figure 44. Checking leaf litterbag at Little River.

are taken to the laboratory for identification, and 

later released into their respective streams.

 What this tells the observer. Leaf-litterbag surveys provide information on: (1) species 

presence (but not absence) at the time of sampling; (2) life-history information, such as larval 

size and activity patterns; and (3) habitat information. Sampling effort is easily quantified.

Limitations. Although the technique 

may be effective for determining the presence of 

many stream-dwelling salamander larvae in 

Great Smoky Mountains National Park, the 

variation in the numbers of individuals captured 

and the inability to relate captures to overall 

abundance make trends impossible to monitor 

without considerable additional effort, such as 

by employing mark-recapture techniques on, 

often, very small larvae. Capture may be influenced by the factors listed in Things to Consider During Planning. Even if every attempt 

is made to standardize sampling (for example, 

by sampling at the same streams during the 

same time of year), environmental factors, as 

well as natural variation in reproductive output, 

likely will be different among years and locations and thus influence whether a species is 

captured. Since environmental and other variables influence the number of animals captured, 

differences in counts over time may not reflect 

changes in status. Additionally, it is difficult to 

determine whether the bags are selected by 

adult and large larval salamanders as places of 

retreat or for foraging, and to determine the 

amount of area actually being sampled using the 

method.

Automated frog call data loggers – Automated data loggers have been used successfully 

to determine the presence of calling frogs at 

breeding sites (fig. 45). They can be set to 

record at variable time intervals for various 

amounts of time throughout the entire day, or 

they can be programed to record only at certain 

times of a 24-hour period, such as from dusk to 

dawn. Frog calls are easily discerned by 

listening to the tapes, and it is sometimes 
59

possible to gain an index of calling intensity, 

provided large choruses are not involved.

Example. At a pond the size of Gum 

Swamp, three data loggers could be installed to 

monitor chorusing frogs: one on the east shore, 

one on the west shore, and one on either the 

north or south shore midway between the other 

two. The program could be set to record for 

5 minutes every hour throughout the day, or for 

5 minutes only from dusk to dawn (the starting 

and ending times would vary with season to 

account for day length). Both sides of the tape 

can be used, thus extending the amount of time 

between tape changes. Data loggers measuring 

water and air temperature, and barometric pressure, could be placed near the call logger to 

account for environmental influences on calling 

activity.

 What this tells the observer. Automated 

frog call data loggers provide information on: 

(1) species presence at the time of sampling 

(species likely to be overlooked during timeconstraint sampling can be recorded with 

Figure 45. Recording data in field as storm 

approaches at Cataloochee Divide.

greater reliability); (2) life history and phenology information, such as when frogs call 

(especially if different species call at different 

times of the day), what environmental influences affect calling; and (3) a relative index of 

the number of males calling.

Limitations. Although species can be 

easily identified, categorizing abundance may 

be very difficult in even moderately sized choruses because of call-overlapping interference. 

It is also often not possible to separate individual callers, allowing the possibility that a single 

calling male could be counted multiple times. 

Since environmental variables influence the 

number of animals calling, differences among 

abundance categories over time may be only 

reflective of differences in environmental conditions during sampling periods. Thus, call 

surveys using automated data loggers must be 

conducted at multiple occasions during the 

potential breeding season. Further, call surveys 

tell nothing about the presence and number of 

females and nonbreeding males, or whether 
60

reproduction was successful. Frog call surveys 

using automated data loggers are best 

implemented where researchers have limited 

access by road (such as along lower Hazel and 

Eagle Creeks) or when rare species are 

suspected.

Whereas automated frog call data loggers 

are relatively easy to assemble (appendix IV), 

they are somewhat expensive (about $350 in 

2002). Unfortunately, there are no computer 

programs currently available that can identify 

calls and categorize abundance by reading the 

tapes. Thus, researchers must listen to tapes and 

manually record the results, a time-consuming, 

tedious exercise. At the Florida Integrated 

Science Center, two observers independently 

listen to the tapes as a measure to reduce and 

quantify observer bias. Automated data loggers 

must be well hidden to reduce theft and vandalism, and this can limit their effectiveness. 

Curious bears have been known to investigate 

and attempt to dismember the data loggers.

Intensive Passive Sampling

Traps (aquatic or terrestrial): funnels, 

bottles, minnow, wire basket – Various types of 

aquatic traps have been used to sample amphibian larvae; on occasion, some of these traps 

have been used to capture adults, such as the 

Common Mudpuppy, in fine wire-mesh basket 

traps. They are all based on the premise that an 

animal entering the trap will be unable to escape 

because it would be difficult to exit through the 

inward-directed funnel opening. However, few 

studies have examined this assumption, and 

unhindered movement into or out of a trap 

(termed trespass) undoubtedly occurs with 

varying degrees of frequency. Minnow traps 

come in wire-mesh, collapsible soft, and plastic 

variations. Wire-mesh minnow traps seem to 

capture the most larvae, whereas plastic-mesh 

traps seem to have the least capture success. A 

drawback to wire-mesh traps is that they cause 

injury to tadpoles, even when checked every 

Various types of aquatic traps have been 

used to sample amphibian larvae....

day, because the animals tend to beat themselves 

against the metal mesh attempting to escape. 

Wire-basket traps are usually larger with larger 

mesh, and are more often used to sample fishes 

and turtles than amphibians. In Florida, a modified crayfish trap with a fine mesh plastic insert 

is used to capture aquatic salamanders (Amphiumas, Sirens) (http://www.fcsc.usgs.gov/posters/Herpetology/Sirens_and_Amphiuma/

sirens_and_amphiuma.html). The trap has not 

been tested specifically to capture amphibians in 

more temperate habitats. Wire-mesh screen funnel traps have been used for both aquatic and 

terrestrial sampling. These traps are placed flush 

with a downed log, rock, or drift fence. As the 

animal enters the trap, it falls to the center and, 

presumably, cannot find its way back out of the 

trap. None of these traps are baited, although 

larvae may attract invertebrate and vertebrate 

(turtles, snakes) predators.

Example. Researchers place 15 wiremesh minnow traps around the perimeter and in 

the center of Big Cove Beaver Pond. Traps are 

spaced at about 5 m apart, secured to a branch to 

prevent loss, and placed in such a manner that 

trapped air-breathing animals have access to 

surface air. Traps are checked daily, perhaps 

even once in the morning and once at night. The 

number of animals caught are recorded by 

species, size, and developmental stage, then 

released. Sampling should only require a few 

days at each location, although a location may 

be trapped more than once per season to capture 

both early and late breeders. Sampling effort is 

easily quantified (number of traps x number of 

days = number of trap days).

 What this tells the observer. Funnel traps 

are used to detect a species= presence, and perhaps to obtain a crude abundance estimate (that 

is, very large numbers of larvae versus very few 

larvae). Counts have little meaning except in 

this context. Funnel trapping is often used during mark-recapture studies, especially if there 

are no known capture biases (that is, trap avoidance or trap happiness). Traps might be useful in 

sampling for rare species.

Limitations. Some types of traps require 

assembly, whereas others can be purchased 

ready-to-use directly from a supplier. They are 
61

subject to vandalism by both wildlife (bears, 

pigs) and people; minnow traps, in particular, 

may be stolen. Trapped animals are vulnerable 

to drowning, predation, and injury, making 

daily checking, preferably in the early morning, 

absolutely essential to minimize mortality. 

Traps capture nontarget organisms, such as 

invertebrates and fish. Even if every attempt is 

made to standardize sampling (for example, by 

sampling at the same exact location and during 

the same time of year), environmental factors 

likely will be different and thus influence 

whether a species is captured. It is very difficult 

to determine any kind of population trend based 

on periodic counts since it is unknown what the 

relationship is between the counts and actual 

abundance. Captures also may be biased by trap 

avoidance or trap happiness (that is, returning to 

a trap again and again because of the availability 

of food or shelter). It may be necessary to conduct a pilot study prior to employing trapping 

methods to determine sampling biases.

Drift fences – Drift fences are the most 

labor intensive method for sampling amphibians. In brief, the idea is to intercept an animal 

during its daily wanderings, direct it along a 

fence constructed of metal (galvanized or aluminum) or cloth (highway department silt cloth; 

plastic sheeting) to where it either falls into a 

pitfall trap (a bucket or can sunk flush with the 

ground surface) or funnel trap (wire-mesh 

screening with inward-directed funnels; once 

the animal gets inside the funnel, it should be 

difficult for it to escape). Sometimes buckets 

and funnels are used simultaneously. There are 

a number of different array configurations, but 

they usually take some form of a Y or X shape; 

each arm is 7.5-10 m long. Drift fences also can 

be used to completely encircle breeding ponds. 

Each sampling unit may consist of three or four 

arrays randomly placed in an area. In a region 

the size of the Great Smokies, dozens of arrays 

would be necessary to sample the terrestrial 

amphibian communities. Arrays should be 

opened at least four times per year for a minimum of 2 weeks per sampling period; at high 

elevations, the winter sampling period could be 

skipped. There are several excellent descriptions of the technique and various configurations, and the reader is referred to chapters in 

Vogt and Hine (1982) and in Heyer and others 

(1994) for more information.

Example. Researchers decide to use a 

Y-shaped drift fence configuration to sample 

lowland, terrestrial amphibians in the Cades 

Cove region. Twenty sampling locations are 

randomly selected, and three arrays are placed 

at each location approximately 50 m from one 

another. The fence must be trenched so that 

animals cannot walk underneath the fence, and 

so that erosion does not create areas for underfence trespass. Pitfalls may not be feasible 

because of the rocky soils, so two funnel traps 

are placed on each side of a fence arm (that is, 

12 per array). Funnel traps may need to be 

shaded to prevent desiccation of trapped animals and are placed flush with the base of the 

fence. Traps must be checked daily to avoid 

animal desiccation and minimize predation. 

The number of captured individuals of each 

species for each funnel trap is recorded. Animals are released at least a few meters away in 

appropriate cover to minimize chances of 

recapture. Funnel traps are opened and checked 

four times per year for a period of 2 weeks per 

sampling occasion to ensure that different 

amphibian faunas are sampled (that is, those 

species which are active during the cool versus 

the warm times of the year). 

 What this tells the observer. Drift fence 

surveys provide information on: (1) species 

presence (but not absence) at the time of 

sampling; (2) life history information, such as 

population size-class structure, reproduction, 

and activity patterns; and (3) when used with 

mark-recapture techniques (toe-clipping, 

elastomer marking, photographic identification), to obtain a measure of abundance. A drift 

fence-pitfall-funnel trapping regimen might be 

useful in capturing rare species or, when completely encircling a breeding site, in measuring 

reproductive effort and success. Sampling effort 

is easily quantified (number of buckets or 

funnels x the number of nights over which the 

sampling was conducted = number of bucket- or 

trap-nights).

Limitations. Drift fences take a great 

deal of work to install and maintain, even without digging holes for pitfalls and carrying heavy 
62

metal flashing to a study site. They are subject 

to vandalism by both wildlife (bears, pigs) and 

people; drift fence materials may also be stolen. 

Animals are very vulnerable in pitfalls and 

traps, making daily checking, preferably in the 

early morning, absolutely essential to minimize 

animal desiccation and predation from reptiles 

and small and large mammals. Pitfalls also capture large numbers of shrews which either eat 

the other animals present or die from stress.

As previously mentioned, the probability 

of catching an animal is influenced by all the 

factors listed in Things to Consider During 

Planning. Even if every attempt is made to 

standardize sampling (for example, by sampling 

during the same time of year), environmental 

factors likely will be different and thus influence whether a species is captured. Since environmental variables influence the number of 

animals that are active, differences in captures 

over time may be more reflective of differences 

...differences in captures over time may be 

more reflective of differences in environmental conditions... 

in environmental conditions among the yearly 

sampling periods than changes in status. It is 

very difficult to determine any kind of trend 

based on periodic counts since it is unknown 

what the relationship is between the counts and 

actual abundance, unless mark-recapture techniques are employed. 

Even with mark-recapture techniques, 

only a very small portion of the population may 

be sampled (for example, terrestrial plethodontids may be territorial and thus unlikely to move 

about very much), so it may be difficult to 

extrapolate estimates of abundance in a wide 

area where animals are patchily distributed. 

Recapture rates are notoriously low in most 

mark-recapture studies of terrestrial salamanders, making estimates of variance quite 

high and unacceptable. Many amphibians may 

not walk along a fence (treefrogs might just 

climb it, hop over, or just pass it by), enter a funnel, or fall into a pitfall; some amphibians may 

be readily able to crawl out of a pitfall. Little is 

known about capture biases, but data from other 

studies indicate that the color (Crawford and 

Kurta, 2000) and size of the bucket may influence capture; that some individuals learn to 

avoid buckets; and, that other individuals may 

come to recognize buckets as a source of shelter 

or food. Therefore, capture probabilities are 

likely to vary considerably among species, even 

if the species is locally abundant.
63

DATA HANDLING

Field Data

Field data should be recorded immediately when taken (fig. 45). Data may be recorded on 

data sheets, preferably in pencil using waterproof paper, or by using preprogrammed palm pilots. 

ARMI is currently developing a web based data entry program using palm pilots. Park researchers 

may desire to link their data collection with the DOI-sponsored national amphibian monitoring 

program. Palm pilot programs with project-specific formats also can be developed. In any case, the 

following data should be recorded at all sampling sites (note that all measurements should be 

recorded in metric units):

Date: month/day/year.

Site No.: a unique identifying site number. 

Example: BB-1 could indicate site 1 on the 

Bunches Bald Quadrangle. There are many 

ways this can be done, but site location codes 

should be consistent.

Personnel: initials or names of those persons 

conducting the survey.

Weather: at the time of the survey.

Altitude: in meters.

Wind: categorical judgement of wind speed 

1 m above sampling area.

General location: a geographic description of 

the site location. Example: Garretts Gap on 

the Hemphill Bald Trail on Cataloochee 

Divide.

Specific location: using GPS or Topo7 software.

Quadrangle: USGS 7.5' quadrangle map.

Start time and End time: in military time (that 

is, 0800 or 1600 hrs).

Standing water: at aquatic sites, record whether 

water is present.

Water level: deepest water level at sampling 

site. Can be estimated (example: > 0.5 m).

Air temperature (AT): recorded at 1 m above 

substrate in EC.

Water temperature (WT): recorded at 30 cm 

under water in EC.

Substrate temperature (ST): recorded at 30 cm 

under leaf litter in EC.

Relative humidity: recorded at 1 m above substrate in EC.

pH: when appropriate, recorded in soil/water 

with a calibrated meter.

Conductivity: when appropriate, recorded in 

water with a calibrated meter.

Habitat type: a general appraisal of the habitat 

type (circle one, see appendix II).

Vegetation: a general appraisal of the 

vegetation types (circle as many as appropriate, see appendix II).

Canopy: a categorical assessment of canopy 

cover (especially important at wetland sites).

Slope aspect: a compass direction of slope 

aspect.

Drainage direction: in which compass direction 

does a stream flow at the sampling location.

Amphibians. The species (using the three letter 

species code), sex (if discernible), life stage 

(adult, juvenile), number of individuals, and 

other notes (for example, reproductive condition, missing limbs) should be recorded. In 

some cases, the snout-vent length (for salamanders), total length (for frogs), mass, or 

other individual measurements may be 

required by a study=s objectives. Measurements should always be in metric units.

Method of capture: specialized capture techniques may require a data form to reflect the 

types of data taken, in addition to the information listed above. For example, the identifying number of the trap, PVC pipe, or 

coverboard should always be recorded to 

discern possible capture biases. The distance 

an animal is captured or observed from a 

transect=s origin and baseline helps indicate 

spatial distribution.

Invertebrates: the type (genus, order, class) and 

relative abundance of invertebrates may be 

very important in studies of amphibians, 

especially amphibians breeding in ponds and 

woodland pools.

Active sampling effort: the number of observers (exclusive of the person recording data, 

unless that person is also sampling animals) 

x the amount of time sampling occurs.
64

If data sheets are used, additional information concerning the site can be included on 

the back of the form, such as drawings of ponds 

or pools, sketches and notes of unusual color 

patterns or morphology, notes on the physical 

description of the sampling site, records of photographs taken, and the presence of unusual 

plants and animals. A sample data sheet is 

included as appendix II.

Spreadsheets and Databases

Most U.S. Government agencies are now 

using Microsoft Excel7 and Access7 to generate spreadsheets and databases. Data from field data 

sheets should be transferred into one of these 

programs as soon as possible following a survey, or 

entered directly while in 

the field using palm pilots, 

using the same conventions as on the data sheets. 

Both programs are compatible with a variety of 

statistical programs, such as SAS7 (Statistical 

Applications Systems). Data accuracy should 

be checked to ensure quality control and prevent 

inaccuracy; the field data sheets serve as a 

backup from which to double check data 

records. Backup copies of data should be made 

weekly, at a minimum, and copies should be 

safely stored at different physical locations or in 

a fireproof data safe.

Analysis and Software

The objective of monitoring the amphibians of Great Smoky Mountains National Park is 

to detect population trends so that actions can be 

taken, if possible, to reverse declines should 

they be detected. Inasmuch as many species=

populations fluctuate from one year to the next, 

especially in unstable habitats such as temporary ponds, and that populations probably go 

extinct naturally (and vacant habitats are recolonized), trend analysis is not an easy task to 

apply to amphibian populations. Much ongoing 

research is focused on amphibian populations; 

new biometric methods are being developed to 

analyze trends in light of the complexities of 

amphibian biology.

Traditionally, population trends have 

been measured via changes in numbers or abundance of the animal in question. If the population size can be measured through time, then 

changes could indicate increasing or decreasing 

trends and, therefore, reflect changes in conservation status. To determine the size of a population, it is necessary to relate the numbers 

recorded during periodic counts to the overall 

population size. The most commonly used 

method to do this is to individually mark animals and to record the numbers recaptured during a period of extended sampling. Thus, each 

animal is accorded a capture history. If enough 

animals are captured and recaptured during a 

survey, it is possible to relate the counts mathematically to an estimate of actual population 

size within a certain degree of confidence. 

Although it is beyond the scope of this manual 

to discuss the nuances, theory, and assumptions 

of mark-recapture analysis, there is substantial 

literature available on this subject (Pollock and 

others, 1990; Nichols, 1992; Thompson and 

others, 1998).

Unfortunately, it is not easy to use markrecapture techniques when studying populations 

of amphibians for two reasons:

1. Amphibians are not easy to mark “permanently.” Various methods, such as toe clipping, elastomer implants, and 

photographic identification (ID), have been 

used, although each technique has limitations. Amphibians lose toes naturally and 

regrow clipped toes; elastomers are time 

consuming to apply and are difficult to read 

under field conditions, and photographic 

ID is not practical when hundreds or thousands of animals are involved or when animals are uniformly patterned or unpatterned. 

Observer error is an ever-present bias.

2. In most instances, very few recaptures are 

recorded in relation to the number of 

amphibians marked. In such cases, the 

variance of the population estimate can 

become quite large, thus negating the reliability of the estimate.

In the Great Smokies, there is only one 

species, the Hellbender, that is probably amenable to reliable mark-recapture population 
65

estimation. These large salamanders are territorial and relatively confined to a circumscribed 

habitat (only large rivers and streams) in a few 

areas of the Park. They can be permanently 

identified through implantation of an injectable 

passive integrated transponder (PIT) tag. As 

such, resurveys should be possible to track populations within certain sections of streams. 

Nickerson and others (2002) have marked 

Hellbenders in Little River using PIT tags, and 

National Park Service biologists should be able 

to track the status and size of this population 

annually using a transect-based snorkeling 

protocol.

Another technique that is gaining favor is 

to conduct repeated sampling at locations 

throughout a designated area, such as a Park or 

refuge, or in a particular subset of a habitat type 

within such an area. Through time, researchers 

can record a capture history for each species at 

each location. Thus, a data set is developed that 

in practice looks very much like the capture history of individuals in a typical mark-recapture 

study. By recording changes in these species=

capture histories through time, biometricians 

can determine detection probabilities for each 

species. Trends can be determined by changes 

in the “percent of area occupied” (PAO) by a 

species and by changes in detection probabilities. More information on applying PAO analyses to monitoring amphibians is contained in 

MacKenzie and others (2002), and at: 

http://www.mbr-pwrc.usgs.gov/

software.html#presence

SOFTWARE

Program MONITOR – Power analysis 

basically tells the researcher how reliable his or 

her data are considering a number of variables, 

such as sample size and the length of time that a 

program is conducted. Important caveats for 

interpreting the results of a monitoring program 

are contained in “Power Analysis of Wildlife 

Monitoring Programs: Exploring the 

Trade-Offs Between Survey Design Variables 

and Sample Size Requirements” by Paige C. 

Eagle, James P. Gibbs, and Sam Droege (http://

www.pwrc.usgs. gov/resshow/droege3rs/

salpower.htm). The USGS has developed a free 

software program, MONITOR, which uses 

linear regression to estimate the statistical 

power of population monitoring programs relative to: the number of plots monitored, the magnitude of counts per plot, count variation, plot 

weighting schemes, the duration of monitoring, 

the interval of monitoring, the strength and 

nature of ongoing population trends, and the 

significance level associated with trend determination. MONITOR is available at:

http://www.mbr-pwrc.usgs.gov/software.html

(then click on POWER)

Program MARK – Program MARK 

provides population parameter estimates (for 

example, survivorship and population rate 

changes) based on mark-recapture data. 

Re-encounters (captures or observations) can be 

recorded from animals found dead, live recaptures (for example, the animal is retrapped or 

resighted), radio tracking of an animal=s movements, or from some combination of these 

sources. The time intervals between re-encounters do not have to be equal, but are assumed to 

be one time unit if not specified (for example, 

every week or month). Data can be subsetted, 

such as by sex or life history stage, so that population parameters can be estimated for the 

designated group. The basic input to program 

MARK is the encounter history for each animal 

(for example, the entry 1001101001 could 

result for an animal caught 5 times during 

10 sampling periods where 1 = captured, 0 = not 

captured). MARK also can be used to provide 

estimates of population size for closed populations. Capture and recapture probabilities for 

closed models can be modeled by attribute 

groups and as a function of time, but not as a 

function of individual-specific covariates. 

Program MARK is available free from 

Colorado State University at:

http://www.cnr.colostate.edu/~gwhite/mark/ 

mark.htm

Program PRESENCE – The number 

and diversity of amphibians in the Great 

Smokies and elsewhere in the southeast makes 

monitoring all species difficult, if not impossible. Nonetheless, high species richness of 

Power analysis basically tells the researcher 

how reliable his or her data are...
66

amphibians is a hallmark of ecosystems in 

southeastern North America. Changes in ecosystems through disturbance, human activities, 

disease, environmental contaminants, or other 

factors could negatively impact the composition 

and richness of amphibian communities. Estimating variation in species richness through 

time and among different locations is one 

means of tracking the status of amphibians as a 

group. This type of analysis, termed percent of 

area occupied (PAO), may be more effective 

than focusing on abundance measures of individual species, which have been shown in most 

studies to lack statistical power because, in part, 

of the low recapture probabilities in markrecapture studies of amphibians.

In the past, the main hindrance to making 

reliable inferences about variation in species 

richness has been the inability to count all species present in an area during a survey. Weather 

conditions, the behavior of different species, 

cryptic coloration, and observer skill are just 

some factors affecting detection (also see 

Things to Consider During Planning). Invariably, some species will be missed, thus biasing 

the estimates (Boulinier and others, 1998a,b). 

However, methods are now available which 

account for variation in detection probabilities, 

and which estimate species richness, standard 

error, and 95 percent confidence intervals 

(Nichols and Conroy, 1996). These methods 

have been extended to estimate several 

important vital rates in animal communities, 

which would be useful to assessing status, for 

example, rates of local species extinction, turnover, and colonization (Nichols and others, 

1998a). They also have been used to test 

hypotheses concerning factors affecting temporal (Boulinier and others, 1998a,b) and spatial 

variation (Nichols and others, 1998b) in species 

richness.

The application of PAO methods to 

amphibian survey data is promising, not only 

because these methods can address important 

questions, but also because they may easily be 

applied to inventory surveys, intensive monitoring at preselected sites, and in extensive surveys 

(MacKenzie and others, 2002). Furthermore, 

detection of a change in species richness can 

alert biologists and managers to potential problems that may require more focused study. To 

facilitate PAO analyses in amphibian monitoring studies, USGS researchers have developed 

Program PRESENCE. This program is available free at: http://www.mbr-pwrc.usgs.

gov/software.html#presence. This program is 

still being tested and developed; undoubtedly 

improvements will be forthcoming to enhance 

its performance and ease of use.

Field researchers require adequate 

equipment and training before undertaking 

amphibian inventory and monitoring activities 

in Great Smoky Mountains National Park. Volunteers can be trained to conduct supervised 

activities, such as call surveys, but quality 

assurance and control must be maintained by a 

supervising biologist. Identifying the amphibians of Great Smoky Mountains National Park is 

often complex and difficult (Dodd, 2004). Even 

experienced herpetologists are sometimes 

unable to verify identification to species, 

especially among salamanders of the genus 

Desmognathus and for many salamander and 

frog larvae (notably very small animals). 

Experienced judgement is critical to a successful monitoring program.

Before going into the field, survey crews 

must be instructed in the proper use of survey 

techniques and map reading, and each crew 

member should be instructed in the use and care 

of each piece of equipment. Prior to beginning 

surveys, field trips should be conducted to 

examine the major amphibian communities, 

and to gain hands-on experience with identification, specifically with regard to key characters. 

Field crews should be taught why certain techniques are being used, the limitations of those 

techniques, and what the results will tell the 

researcher. Communication is important to 

minimize observer bias, a major cause of error 

in field studies. Individuals should be made to 

feel part of the team, and they should be 

credited for hard work under sometimes 

EQUIPMENT AND TRAINING
67

difficult conditions, as well as for the discoveries made.

To assist planning, a checklist is provided 

in appendix III for equipment needed at field 

sites during amphibian surveys and data collection. All crews should be briefed on the dangers 

of hypothermia, heat stress, lightning, and 

dangerous animals (yellowjackets and wasps, 

venomous snakes, pigs, bears, humans). Each 

vehicle should have appropriate first aid, safety, 

and communications supplies. Crews should be 

properly dressed for cold or heat and inclement 

weather, especially with regard to footwear. 

Never conduct surveys, even in streams, in bare 

feet or sandals because of the dangers of sharp 

rocks or glass. Crews should always provide a 

destination and estimated time of return to 

supervisors before setting out on surveys.

Concern about disease and toxic contamination as causes of amphibian declines 

has increased considerably in recent years 

(Carey and Bryant, 1995; Daszak and others, 

1999). A corollary of this concern is the need 

for field workers to avoid becoming vectors 

for transmitting disease organisms or toxic 

chemicals to and among study sites. The 

Declining Amphibian Populations Task Force 

(DAPTF) has developed a standard protocol 

for use by anyone conducting fieldwork at 

amphibian breeding sites or in other aquatic 

habitats. These procedures should be used for 

all routine surveys, but more stringent measures are necessary in areas with known 

diseases.

BIOSECURITY AND DISEASE

Biosecurity Protocol

Protective Wear & Equipment Disinfecting & Sanitizing Methods

nonpermeable boots or waders rinse in bleach solution immediately after leaving each study site3

(fig. 46)

vinyl gloves1 dispose of gloves after each handling incident

nets rinse in bleach solution immediately after leaving each study site

plastic bags (for holding specimens)2 properly dispose after each use

needles & syringes (for blood extraction) properly dispose after each use

scalpel blades, PIT tag cannula, forceps, etc. immerse in sterilizing solution

1

Only vinyl gloves should be used when handling amphibians. Some people are allergic to latex gloves, and latex gloves are toxic

to amphibians (Gutleb and others, 2001). 2

Use one bag per specimen.

3

Premixed bleach solutions can be carried in containers large enough to step into and immerse boots, nets, and equipment. If this is 

not possible, bleach solutions can be carried in a spray backpack firefighting pump.

Solution Formulas

bleach one (1) capful per gallon water

sanitizing solution (for instruments) 70% methanol for 30 minutes, then flamed; or, 1% glutaraldehyde for 15 minutes; 

or, boiling water for 10 minutes
68

Additional Precautions

X Avoid contact between used and unused 

protective wear and equipment.

X Separately house specimens.

X Avoid contact between gloved hands and 

face, especially the area of the nose.

X Do not urinate in or near ponds and 

streams.

X Wash hands thoroughly with soap and 

water, or use a sanitary wipe, after urinating.

X Wash hands thoroughly with soap and 

water, or use a sanitary wipe, after handling specimens known or suspected of 

being diseased or contaminated.

X Wash hands thoroughly with soap and 

water, or use a sanitary wipe, after leaving 

each site.

X Do not use insect repellent on hands when 

handling amphibians.

Figure 46. Biosecurity. Washing boots and 

stump ripper in bleach solution.

Disease Protocols

The following information is taken from 

the U.S. Geological Survey=s STANDARD 

OPERATING PROCEDURE (Kathryn Converse and D. Earl Green; ARMI SOP No. 105; 

revised March 2, 2001) entitled “Collection, 

Preservation & Mailing of Amphibians for 

Diagnostic Examinations.” It was developed by 

the National Wildlife Health Center, Madison, 

Wisconsin (http://www.nwhc.usgs.gov/

research/amph_dc/sop_mailing.html).

The best diagnostic specimen is the live, 

sick amphibian. Live amphibians are necessary 

to obtain meaningful bacterial cultures and most 

types of fungus cultures. In addition, blood for 

various “blood tests” can be obtained only from 

live amphibians. Dead amphibians have limited 

usefulness because aquatic animals decompose 

much more rapidly than terrestrial animals 

which means amphibian carcasses nearly 

always will have large numbers of decompositional bacteria and fungi throughout their bodies. This rapid decomposition (autolysis) makes 

it very difficult to obtain meaningful or useful 

bacterial and fungal cultures, but dead amphibians may still have usefulness for virus cultures, 

histology and toxicological tests, if promptly 

and properly preserved.

If the amphibians will be captured and 

euthanized as part of other studies, then first 

observe and record their behavior. Blood should 

be collected and saved prior to euthanasia. If the 

euthanized amphibians will be preserved in a 

fixative, then collect swabs for bacterial, viral 

and fungus cultures from the mouth, vent, skin, 

and any skin abnormalities (lesions) prior to 

emersion of the animal in the fixative. 

At a casualty site, the priority specimens 

for diagnostic examinations are live, sick 

amphibians. Divide dead amphibians into two 

groups: promptly preserve about half the carcasses (preferably the most recently dead 

amphibians) in 10 percent formalin (or 

70-75 percent ethanol); promptly freeze the 

other dead amphibians (for virus cultures and 

possible poison tests). In cases involving less 

well known species, submission of live healthy 

amphibians as “control” or “baseline” specimens will be necessary to assist in the interpretation of findings in the sick or dead animals. 
69

More than one lethal disease may affect a population simultaneously, so submission of multiple animals is always encouraged. Collect 

specimens that represent the species that are 

affected and the geographic areas. Do not place 

live and dead animals in the same container, and 

do not put multiple species in the same container (except, it is acceptable to put dead 

animals of multiple species in one container of 

formalin or ethanol).

If possible, submission of invading (alien 

or introduced) amphibians from the casualty 

site is desirable, even if they appear healthy or 

unaffected, because invasive species can be the 

vectors of infectious diseases. If any other 

endemic amphibians, fish, or reptiles are 

present at the casualty site, these animals also 

may need to be examined as part of a wider 

epizootiologic investigation into the cause of 

the casualties. 

Many amphibian die-offs are fleeting. 

This means the casualties must be collected the 

hour and day they are found. Returning to the 

casualty site the next day to collect sick 

amphibians and carcasses invariably fails 

because of the highly efficient activity of scavengers during the night and rapid autolysis of 

carcasses.

LIVE AND SICK AMPHIBIANS

Eggs – Place eggs in heavy mil plastic bag or 

plastic container. Equal volumes of air and 

water should be present in the bag or container to assure adequate oxygen exchange. 

Do NOT fill bags or containers completely 

with water. If bottled oxygen is available, it 

may be placed into the air cell in the bag or 

container, but this is optional. If possible, 

place plastic bags in a solid container for 

support and to avoid crushing specimens or 

puncture of the bag. 

Tadpoles, Larvae, and Neotenes – Same as 

for eggs. For small amphibians (<2 grams 

each), multiple live animals may be placed 

in one container, but avoid mixing species. 

For larger aquatic larvae and neotenes, one 

animal per bag or container is recommended. Enough air must be present in 

each container; containers that have a large 

surface area of water to air are preferred; 

hence, flat food storage-type plastic boxes 

with lids (available at nearly any grocery 

store) are preferred to tall narrow plastic 

bottles. If bottled oxygen is available, oxygen may be placed into the air cell in the 

bag or container, but this is optional.

Adult Amphibians (Terrestrial 

Amphibians) – Plastic boxes or bottles 

with wide lids may be used for mailing. 

Sick amphibians should be mailed in 

separate containers. Two or more live adult 

amphibians of the same species may be 

placed in one container, but avoid crowding. Note: if an infectious disease is the 

cause of the casualties, the disease may be 

transmitted between amphibians in the container if more than one animal is placed in 

each container. Wet unbleached (brown) 

paper towels or wet local vegetation should 

be added to the container to prevent dehydration of the animal; do not use sponges, 

because many contain chemicals that are 

toxic to amphibians. Three or more small 

holes should be made in the lid of each 

container. Plastic bags are not recommended for terrestrial amphibians.

DEAD AMPHIBIANS

About half the dead amphibians should be 

immediately placed into 10 percent buffered neutral formalin or 75 percent ethanol 

for histologic examinations. When 

possible, the freshest carcasses (those with 

the least amount of decomposition) should 

be selected for fixation. Prior to immersing 

the carcass in the fixative, slit open the 

body cavity along the ventral midline to 

assure rapid fixation of internal organs. For 

the first 3-4 days of fixation, the volume of 

METHODS
70

fixative to volume of carcasses should be 

10:1. After 3-4 days of fixation, the carcasses may be transferred to a minimal 

amount of fresh fixative that prevents 

drying of the specimen. 

Freezing – About half the carcasses should be 

promptly frozen. Preferred freezing temperature is -40 degrees, but any freezing 

temperature is preferable to a chilled carcass. Do NOT freeze amphibians in water. 

Frozen carcasses can be used for virus cultures, toxicological examinations, and 

molecular (DNA) tests. Frozen and preserved carcasses are not suitable for bacterial and fungus cultures; generally, 

bacterial and fungus cultures will be 

attempted only on amphibians that are submitted live. 

Decomposed Carcasses – Clearly decomposed 

carcasses may have some diagnostic usefulness for molecular testing and toxicological analyses. Very decomposed carcasses 

with fluffy growths of fungus on the skin; 

maggots in the mouth, vent, and body 

cavity; or carcasses of only skin and bones 

should be frozen and saved if fresher carcasses are not available. 

LABELS

Each container must be labeled. Paper 

labels written in pencil are preferred, especially 

if there is ethanol in any containers. Most ink 

will dissolve in ethanol or become streaked 

during freezing and thawing. Each label should 

have the following information:

H species

H date collected

H location (state/county/town)

H found dead or euthanized

H collector (name/address/phone)

H additional history on back of tag

MAILING

Shipping Container – Use a picnic cooler or 

styrofoam-lined cardboard box. 

Ice – Ice packs (blue ice) is preferred to wet ice 

to avoid leaking during shipment. Most 

amphibians from temperate climatic zones 

should be mailed with ice packs. Ice packs 

should be wrapped with about 5 layers of 

newspaper before being placed at the side 

of containers of amphibians. For live 

amphibians, position ice packs on the side 

of the shipping container, not under the 

specimens, as this allows live amphibians to 

move away from cold zones. 

Frozen Specimens – Frozen samples should be 

mailed with dry ice. Ice packs are an alternative, especially if the ice packs were frozen in an ultra-low freezer (-40 or lower). 

Avoid mailing frozen specimens in the 

same shipping container as live animals or 

specimens in formalin. If frozen samples 

and live amphibians (or specimens in formalin) must be mailed in the same shipping 

container, never put dry ice in the shipping 

container. If frozen samples and live 

amphibians (or specimens in formalin) 

must be mailed in the same shipping container, separate the shipping container into 

two compartments with styrofoam panels 

and place the ice packs at one end of the 

container next to the frozen samples. 

Preserved Specimens – Once specimens have 

fixed in a large volume of formalin or ethanol for 3-4 days, the preserved samples may 

be mailed in a minimal amount of preservative that prevents drying. It is not necessary 

to mail large volumes of liquid fixative. 

Preserved carcasses may be wrapped in 

gauze or a paper towel that is moistened 

with the fixative. If preserved specimens 

are transferred to plastic bags, always 

double-bag the specimen and pack it into 

the shipping box to avoid crushing the 

sample during transport. 

Packing the Shipping Container – Plastic 

boxes and bags containing live amphibians 

may be stacked, but keep air holes clear; 

some plastic boxes will stack tightly on 

each other and may seal air holes of lower 

containers. Do not place live amphibians 

directly on top of ice packs, because this 

may cause water in the animal's container 

to freeze. After placing ice packs and specimen containers in the shipping box, add 

crumpled newspaper, plastic peanuts, or 

other filler around the containers to 
71

minimize shifting of contents during mailing and crushing the plastic-bag samples. If 

a styrofoam-lined cardboard box is being 

used for mailing, then line the box with a 

heavy mil plastic bag and place all ice 

packs and specimens into the bag to minimize leaks and moisture condensation into 

the cardboard box. 

Double Bagging – Frozen samples and specimens in formalin (or ethanol) should be 

double bagged. This is especially important 

to avoid fixative leakage. If glass vials or 

jars must be mailed, these too should be 

placed into a plastic bag. 

Taping – Tape should be wrapped completely 

across the lid, sides, and bottom of each 

plastic cooler in at least two places to prevent accidental opening of the container 

during mailing. Nylon-reinforced tape is 

recommended, but 2-inch-wide clear tape 

also may be used. 

Overnight Couriers should be used for sick, 

live, and frozen amphibians. 

Dates for Mailing – Only mail boxes of specimens by overnight couriers on Mondays, 

Tuesdays, and Wednesdays. Most diagnostic laboratories are not open on weekends, 

so specimens mailed on Fridays may be 

held in hot or freezing delivery vans over 

the weekend. A significant percentage of 

packages mailed by overnight courier on 

Thursdays, do not arrive in 24 hrs, and 

these can suffer the same fate. 

Mailing – Overnight courier service should be 

used. Securely tape the cooler or box and 

mail to: National Wildlife Health Center, 

6006 Schroeder Road, Madison, WI 53711. 

Note: in addition to the NWHC address, 

add DIAGNOSTIC SPECIMENS--WILDLIFE to the outside of the box. This label 

will direct coolers with specimens to our 

necropsy entrance. Do not label the 

container with statements like, “Live Animals,” as this could interrupt or prohibit 

shipment because of courier policy. Contact NWHC (608-270-2400) (FAX 

608-270-2415) prior to shipping animals 

by 1 day (overnight) service and after shipment to confirm the estimated time of 

arrival.

QUARANTINE OF AMPHIBIANS

Amphibians (dead or alive) from a casualty site should be considered contagious specimens. Live, sick animals and carcasses should 

never be released or discarded at other sites and 

should not be taken into laboratory settings with 

other live amphibians, fish, or reptiles. Release 

of sick amphibians or discarding carcasses at 

other sites may result in the spread of infectious 

diseases.

In certain parts of North America, 

particularly in the Midwest and northern New 

England, large numbers of malformed amphibians have been observed. Malformations 

involve missing or supernumerary digits, arms, 

or legs, missing eyes, and deformed jaws 

(Meteyer, 2000). Several hypotheses have been 

tested as causes, including parasite-induction 

during development (Morrell, 1999; Johnson 

and others, 2002), the effects of toxic chemicals 

(pesticides), and high levels of UV light; all 

have induced malformations under laboratory 

and field conditions. As with other environmental influences, however, it is possible that the 

malformations observed result from interactive 

causes. Much research is being directed toward 

understanding amphibian malformations.

Fortunately, no malformations of 

amphibians have been found in Great Smoky 

Mountains National Park. The U.S. Geological 

Survey has developed a standardized protocol 

for reporting and handling malformed amphibians (http://www.npwrc.usgs.gov/narcam/

index.htm); should such individuals be found 

within the Park, these protocols should be 

followed.

MALFORMATIONS
72

CONCLUSIONS

concepts, problems, considerations, and 

approaches were outlined for establishing a 

monitoring program for the amphibians of 

Great Smoky Mountains National Park. The 

monitoring approach that is selected (which 

species will be monitored, where they will be 

monitored, how many sites will be monitored, 

and which techniques will be used) will be 

Litterbags:

Consider litterbags

to increase effort

Stream Sites

Randomly select 30 streams

N=30

High Elevation Sites

Randomly select 5 sites on 20

terrestrial trails; N=100*

High Elevation Sites

Randomly select 5 sites on 15

terrestrial trails; N=75*

High Elevation Sites

Randomly select 5 sites on 10

terrestrial trails; N=50*

Low/Medium Elevation Sites

Randomly select 5 sites on 10

terrestrial trails; N=50*

Low/Medium Elevation Sites

Randomly select 5 sites on 20

terrestrial trails; N=100*

Add Minimum funding sites and:

Little River, Cataloochee, and Big Cove

Add Medium funding sites and:

Bone Valley and Cane Creek

Decision Path for Monitoring Amphibians at Great Smoky Mountains National Park

Whiteoak Sink

Monitor Gourley Pond (for disease)

Minimum Amount of Funding

Monitor Hellbenders at Little River

Same

Medium Amount of Funding

Coverboards

1-2 times per month

at selected sites

Monitor Caves

2 visits per year (spring/fall)

Same

Money Received for Monitoring

Monitor Cades Cove and Finley Cane

Wetlands, Sugarlands, and The Sinks

Site Visits,

Call Surveys

Maximum Amount of Funding

Site Visits,

Call Surveys

Site Visits,

Call Surveys

Low/Medium Elevation Sites

Randomly select 5 sites on 15

terrestrial trails; N=75*

Stream Sites

Randomly select 20 streams

N=20

Stream Sites

Randomly select 35 streams

N=35

Litterbags:

Consider litterbags

to increase effort

Monitor All Hellbender Sites

Whiteoak Sink Whiteoak Sink

Litterbags:

Consider litterbags

to increase effort

Coverboards

1-2 times per month

at selected sites

DECREASING PRIORTY

*N is the total number of sites

Figure 47. Decision path for helping design an amphibian monitoring program at Great Smoky Mountains National Park 

based on three levels of funding (see Conclusion).

determined by the funding (and personnel) 

available and the specific objectives of Park 

managers. In this regard, a three-pronged 

approach to amphibian monitoring within the 

Park is presented in figure 47. The decision path 

is based on minimum, medium, and maximum 

levels of funding, although exact amounts are 

deliberately not specified.
73

Minimum Funding

1. In this and all tiers, Gourley Pond must be 

visited several times a year to monitor the 

effects of disease. 

2. A minimum of two to three visits per year is 

specified for the Park=s most critical 

wetlands. Three of the wetland sites (the 

Finley-Cane ponds, Sugarlands, the Sinks) 

are readily accessible by road; all of the sites 

in Cades Cove (Gum Swamp, Methodist 

Church Pond, Shields Pond, Stupkas Sinkhole Pond, Abrams Creek pools) could be 

visited easily in a single day. Nighttime call 

surveys would greatly increase the efficiency 

of wetland surveys in Cades Cove and elsewhere.

3. Time-constrained techniques could be used 

at the terrestrial and stream sites. If five sites 

could be visited per day, sampling these sites 

would take a two or three-person crew about 

3 weeks to complete the data collection. 

Whiteoak Sink is singled out for sampling 

because of the presence of the Southern Zigzag Salamander (Plethodon ventralis) and 

because of all the readily accessible cave 

openings. Litterbags set early in the year 

could be checked easily throughout the season and thus record species that may be not 

encountered during stream time constraint 

sampling.

Medium Funding

1. In addition to the work considered above, the 

number of terrestrial and stream sampling 

sites could be increased.

2. Hellbenders should be monitored annually in 

the Little River.

Maximum Funding

1. In addition to the work considered above, the 

number of terrestrial and stream sampling 

sites could be increased further.

2. Coverboards could be used to increase longterm sampling effort at selected sites; they 

should be checked once or twice monthly.

3. Hellbenders should be monitored annually at 

all known locations in the Park.

4. Selected caves (Gregorys, Stupkas, the two 

Calf caves) should be surveyed thoroughly 

two or three times a year; other caves should 

be visited, especially in Whiteoak Sink, and 

the openings around the entrance and twilight zones searched for salamanders and 

frogs.

To increase sample size, the same terrestrial and stream sites need not be searched annually. For example, 50 terrestrial sites could be 

searched one year; a second 50 searched the 

second year; and a third 50 searched the third 

year, after which the cycle could be repeated. 

Unfortunately, however, there is a tradeoff with 

this approach. If a rotation is used, sample size 

is increased (a good thing), but the amount of 

time it takes to complete a cycle is greatly 

extended (12 years to get four samples per location). Amphibian populations may change dramatically in this amount of time, and trends 

could be missed or misinterpreted. 

A rotating schedule could also be used to 

vary survey species or areas. For example, 

researchers might decide to alternate Hellbender and cave surveys every other year if 

money became limited. Or, Hellbenders could 

be monitored for 2 years in Little River, and at 

the other locations every third year. Planning is 

absolutely essential, and figure 47 is meant as a 

guide to approaches that might be considered 

rather than an absolute schedule.
74

In many regions of the world, amphibian 

species have inexplicably declined or disappeared, and serious malformations have been 

observed, particularly in the upper Midwest 

region of North America. Causes for the 

declines and malformations probably are varied 

and may not even be related. The seemingly 

sudden declines in many amphibians, however, 

suggests that a vigilant approach is necessary to 

monitor populations and to identify causes 

when declines or malformations are discovered.

In the United States, amphibian declines 

frequently have occurred in protected areas 

which should provide an ideal habitat against 

the most common causes of decline, habitat loss 

and changes in land use. In particular, declines 

in western National Parks have concerned biologists, resource managers, and legislators to the 

extent that Congress authorized the U.S. 

Geological Survey to set up a national amphibian monitoring program on Federal lands to 

develop the sampling techniques and biometrical analyses necessary to determine status and 

trends, as well as identify possible causes of 

amphibian declines and malformations when 

they are discovered.

Great Smoky Mountains National Park is 

the most visited park in the National Park Service system. It is also a center of salamander 

diversity in North America (with 31 species 

recorded historically) and contains a moderate 

number of frog species (13 species recorded 

historically). Because of this diversity, the Park 

was selected as a prototype amphibian monitoring location, and USGS biologists conducted 

intensive sampling throughout all regions and 

habitats from 1998 to 2001. This report presents 

the results of this intensive sampling, beginning 

with an overview of the Park=s amphibians, the 

factors affecting their distribution, a review of 

important areas of biodiversity (particularly 

Cades Cove and the Cane Creek drainage), and 

a summary of amphibian life history in the 

southern Appalachians; it concludes with an 

extensive list of references for inventorying and 

monitoring amphibians.

As part of the project, a variety of inventory, sampling, and monitoring techniques were 

employed and tested. These included widescale visual encounter surveys of amphibians at 

terrestrial and aquatic sites, intensive monitoring of selected plots, randomly placed smallgrid plot sampling, leaf-litterbag sampling in 

streams, monitoring nesting females of selected 

species, call surveys, and monitoring specialized habitats, such as caves. Coupled with information derived from amphibian surveys on 

Federal lands using various other techniques 

(automated frog call data loggers, PVC pipes, 

drift fences, terrestrial and aquatic traps), an 

amphibian monitoring program was designed to 

best meet the needs of biologists and natural 

resource managers within the Park after taking 

into consideration the logistics, terrain, and life 

histories of the species found within the 

2,071 km2

 area of the Park. Each monitoring 

technique was described, including an example 

of how the technique was set up, what the 

results tell the observer, and limitations of the 

technique and the data derived from it.

Survey and monitoring projects are both 

time and labor intensive, and resource managers 

must make the best use of the resources available. For this reason, labor-intensive techniques, such as the use of drift fences with or 

without pitfall traps, and various types of trapping techniques which require continuous 

checking, are not recommended. Because only 

one species of frog (Cope’s Gray Treefrog, Hyla 

chrysoscelis) is likely to be attracted to PVC 

pipe (as a hideaway), PVC is not recommended, 

particularly when the species of frog can more 

easily be detected by listening for calls or by 

employing automated frog call data loggers 

(AFCDL). AFCDL are effective at detecting 

frogs within Great Smoky Mountains National 

Park, but are best employed in areas with extensive wetlands, such as ponds within Cades 

Cove. An extensive guide is included as an 

appendix to this manual with instructions on the 

construction and deployment of AFCDL. 

Coverboards are not recommended because of 

SUMMARY
75

potential biases (in which species and age 

classes are observed) associated with sampling.

Extensive use of both small (10 x 10 m) 

and large (30 x 40 m) plots, either randomly 

sampled or “permanently” established, suggested that plot surveys are inefficient when 

compared with visual encounter (or time constraint) surveys. In addition, it is difficult to 

extrapolate counts obtained during plot surveys 

to actual amphibian abundance, despite efforts 

to standardize survey techniques, locations, and 

timing. Inasmuch as capture-recapture protocols are labor and time intensive, and that recapture rates are usually very low, capturerecapture surveys also are not recommended to 

park personnel.

The most consistent and effective survey 

technique to monitor amphibians within the 

Park, especially considering temporal, personnel, and logistic constraints, is to use visual 

encounter surveys based on repeated site visits. 

The use of leaf litterbags is also an effective 

nondestructive technique for determining the 

presence of secretive salamander larvae in 

streams. Data on presence (present/not 

detected), rather than abundance, is used to 

record a capture history for each species at each 

location. Thus, a data set is developed that, in 

practice, looks very much like the capture history of individuals in a typical capture-recapture study. By recording changes in these 

species= capture histories through time, biologists can determine detection probabilities for 

each species. Trends can be determined by 

changes in the percentage of area occupied by a 

species and by changes in detection probabilities. URLs for free, downloadable software are 

included in this report.

Amphibians in the Park should be monitored in a three-tiered approach, which will 

depend on the amount of funding available. 

With minimum funding, biologists should:

• Monitor Gourley Pond, where disease has 

been reported in the past,

• Monitor other Cades Cove wetlands and a 

few other wetlands with easy access,

• Use call surveys to record presence,

• Monitor 50 high-elevation sites (5 sites 

per each of 10 trails),

• Monitor 50 medium- to low-elevation 

sites (5 sites per each of 10 trails),

• Monitor amphibians in Whiteoak Sink, 

and,

• Monitor sites at 20 randomly selected 

streams (employ litterbags to increase 

sampling effort).

As funding levels increase, the number of 

sites monitored could be increased and species 

with specific habitat requirements (Hellbenders, Cryptobranchus alleganiensis; cave species) can be included. In all cases, visual 

encounter (or time constraint) survey techniques are recommended.

Because disease agents were found 

within the Park (iridovirus and fungus in several 

species at Cades Cove), biosecurity protocols 

must be employed after sampling each wetland 

within this region. All nets, boots, and equipment must be cleansed using a 10 percent 

bleach solution, and researchers should carry 

materials into the field which will allow them to 

process dead, dying, or live amphibians. Disease protocols and instructions for handling 

amphibians suspected of harboring disease 

were developed by the USGS National Wildlife 

Health Center, and are reprinted in this report.

Sampling a diverse amphibian assemblage in an area as large as Great Smoky Mountains National Park, and with limited physical 

access, is not an easy task. Randomization of 

sampling sites is not strictly possible, so some 

form of a stratified sampling paradigm must be 

employed. Depending on the amphibian species 

or community sampled, biologists must use 

trails, watersheds, hydrological units, elevation, 

or other parameters to narrow sampling focus. 

Ultimately, however, rarer species or those with 

specialized habits could be overlooked. Species 

identification also is challenging, and the use of 

experienced survey personnel is critical for 

obtaining factual data. In this regard, USGS and 

Park biologists must establish cooperative 

efforts and training to ensure that the congressionally mandated amphibian surveys are performed in a statistically rigorous and 

biologically meaningful manner, and that 

amphibian populations on Federal lands are 

monitored to ensure their long-term survival.
76

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80
81

Appendixes I, II, and III
82

Appendix I. Location of selected wetland sampling sites

[Locations shown in latitude and longitude]

Ponds

Big Cove Beaver Pond 35 30 27N 83 18 02W

Bone Valley Beaver Pond 35 31 07N 83 40 51W

Finley-Cane Sinkhole Ponds (4) 35 36 37N 83 44 38W

Gourley Pond 35 35 36N 83 47 19W

Gum Swamp 35 35 21N 83 50 17W

Methodist Church Pond 35 36 24N 83 49 01W

Sewage Treatment Pond 35 36 14N 83 46 57W

Shields Pond 35 35 33N 83 48 54W

Stupkas Sinkhole Pond 35 35 23N 83 50 52W

Swampy and mucky wetlands

Cataloochee 35 37 44N 83 06 00W

Cataloochee Trout 35 39 12N 83 04 28W

Indian Creek 35 28 51N 83 24 51W

Little Cataloochee 35 39 43N 83 05 55W

Smokemont 35 33 06N 83 18 35W

The Sinks 35 40 10N 83 39 38W

Woodland pools

Abrams Creek 35 35 40N 83 50 41W

Big Cove Pool 35 30 29N 83 18 02W

Cane Creek 35 39 01N 83 53 15W

35 39 42N 83 52 41W

35 39 07N 83 53 05W

Gourley Sinkhole 35 35 34N 83 47 14W

Sugarlands 35 41 11N 83 32 17W

Tremont Roadside Ditches 35 39 15N 83 42 08W

35 39 07N 83 41 47W

Grassy pools (Cades Cove)

35 36 19N 83 47 44W

35 36 20N 83 48 31W

35 36 03N 83 48 33W

35 36 16N 83 48 10W
83

Appendix II. Example of a field data sheet prepared for amphibian surveys in Great Smoky Mountains National Park.

GREAT SMOKY MOUNTAINS NATIONAL PARK

AMPHIBIAN SURVEY FORM

Date:_________________ Site No. __________ Personnel ____________________________

Weather: clear / partly cloudy / cloudy / rain / fog / other:_ ___________________________________

Altitude:_____________ft/m Wind: calm / slight breeze / moderate / windy

General Location: _ _________________________________________________________________

Specific Location (UTM): E _______________; N _______________; Quad: _ __________________

Start Time: ___________ Standing Water: Y / N Water Level: ______________ m

End Time: _________ AT: _____; WT: ______; ST: ______; RH: ______; pH_____; Cond _ _____

Habitat Type (circle one): terrestrial / large stream / med. stream/ small stream / seep / pond / woodland 

pool / mucky area / open grassy pools / cave / rock face / other: ____ ____________________________

Vegetation: spruce-fir / deciduous / cove hardwood / oak / pine / hemlock / open field / other: _______

Canopy: open / sparsely covered / closed Slope Aspect:___________ Drainage Dir: _ ___________

Method of capture: tc= time constraint; ac= area constraint; em= egg mass count; nets= dip or sweep nets;

PVC= pvc pipe; cb= cover board; ft= funnel trap; FL=frog logger; mt= minnow trap; pf= pitfall; tr= transect; 

V= visual; C= calling. Life Stage: A= adult, SA= subadult; L= larvae, E= eggs.

Invertebrates present: Y / N Species:___________________________________________________________

Active Sampling Effort: _____________________________________________________________________

Further notes should be written on back of sheet

Amphibian

Species

No. of

Individuals

Life

Stage

Method of

Capture Notes
84

Appendix III. Sampling equipment. Trademark names are mentioned for information purposes only, not as 

an endorsement of the U.S. Geological Survey.

On the Trail

Weatherproof flat map case containing:

USGS 7.5' topographic maps

Great Smoky Mountains National Park trail

guide

Field Data Sheets (on Rite in the Rain7

paper)

Notebook

Pencils/sharpener

Copy of permits

Equipment (in a sealable rainproof bag):

GPS

Compass

Clinometer

Palm pilot (where appropriate for data entry)

Digital temperature gauge

Extra temperature gauge probe

Digital relative humidity meter

Camera, preferably digital

Binoculars

Pesola7 spring scales (10 g, 50 g, 100 g)

Ziploc7 bags for carrying and weighing

animals (various sizes)

Clear plastic metric ruler

Small mesh hand dip net (for larvae)

Hand lens

Stump ripper (Fuhrman7 Diversified)

Extra batteries

Small pen light (for searching crevices)

Leatherman7 tool

Fluorescent flagging

Biosafety:

Ziploc7 bags for dead or diseased animals

(various sizes)

Sanitary Handwipes

Vinyl gloves (several pair)

Other:

First-aid supplies

Sting-eze7 or other sting remedy, especially

if anyone is allergic to yellowjackets

Extra water

Lightweight cell phone (may not work in

valleys or remote areas)

At Ponds (in addition to the above)

Oxygen, pH, Conductivity meters

Dip nets (both large and small of appropriate

mesh size to capture larvae)

Waders

Shrimper (Lacrosse7 Majesty 12") boots

(excellent in small marshes and creeks)

Tape measure (50 or 100 m) or laser range

finder

Meter ruler

Biosafety:

Pre-mixed bleach solution in appropriatelysized tub or in a spray backpack firefighting pump
Appendix IV

Guidelines for building and operating remote field recorders 

(automated frog call data loggers)

by

William J. Barichivich

87

Guidelines for building and operating remote field 

recorders

(automated frog call data loggers)

By William J. Barichivich1

Automated frog call data loggers have 

been used successfully to provide information 

on: (1) species presence at the time of sampling 

(that is, species likely to be overlooked during 

time-constraint sampling can be recorded with 

greater reliability); (2) life history and phenology information, such as when frogs call (especially if different species call at different times 

of the day), what environmental influences 

affect calling; and (3) a relative index of the 

number of males calling. Although species can 

be easily identified, categorizing abundance 

may be very difficult in even moderately sized 

choruses because of call-overlapping interference. It is also often not possible to separate 

individual callers, allowing the possibility that a 

single calling male could be counted multiple 

times. Because environmental variables influence the number of animals calling, differences 

among abundance categories over time may be 

only reflective of differences in environmental 

conditions during sampling periods. Thus, call 

surveys using automated frog call data loggers 

must be conducted at multiple occasions during 

the potential breeding season. Further, call surveys tell nothing about the presence and number 

of females and non-breeding males, or whether 

reproduction was successful. Frog call surveys 

using automated data loggers are best implemented where researchers have limited access 

by road or when rare species are suspected.

1Florida Integrated Science Center

7920 N.W. 71st Street

Gainesville, Florida 32653

jamie_barichivich@usgs.gov

What is an automated frog call data 

logger?

Automated frog call data loggers are 

recorders that can be programmed to operate for 

a specified duration at specified intervals (for 

example, one minute every hour) and over a 

specified period (for example, 18:00 until 

06:00). They can operate remotely without 

maintenance for extended periods under most 

environmental conditions, including extreme 

heat, cold, rain, and snow. The automated frog 

call data logger described in this section is a conglomeration of stand-alone components, 

whereas the original design (Peterson and Dorcas 1992, 1994) required building several components on a printed circuit board or using an 

expensive commercial data logger to control the 

tape recorder. The literature regarding previous 

designs is helpful and should be reviewed not 

just for construction details but also for study 

design (see related literature).

Why build an automated frog call 

data logger?

Automated frog call data loggers produce 

an archivable record that can be analyzed or confirmed at a later date. Unlike standard aural surveys, no observers are present, so the behavior of 

calling anurans is likely unaffected. Since automated frog call data loggers can be deployed 

prior to monitoring, they can synchronously 

monitor any number of sites 24-hrs/day.
88

How do you build an automated frog 

call data logger?

The basic automated frog call data logger 

consists of an analog tape recorder, timer(s), 

power source (battery) and voltage step-down, 

container, and microphone (figs. 1-2). The battery powers the timer(s) that regulate power, 

also from the battery, to the tape recorder. The 

tape recorder, timer(s), battery and voltage stepdown are housed in a weatherproof container 

and an external microphone picks up nearby 

sounds (for example, frog calls) and relays the 

signals to the tape recorder in the container 

(fig. 1).

Construction

1. Parts-gathering materials can be time consuming. Vendors are often out of stock and 

no single source carries all the necessary 

components to build an automated frog call 

data logger (table 1).

a. A wide range of analog tape recorders

have been utilized and can range in price 

from tens to hundreds of dollars. Since 

the recorder is the heart of the system, 

consider the highest quality recorder 

within reason. The following features are 

highly desirable:

(1.) Stereo recording provides left and 

right channel recording.

(2.) Extended record time slows the 

speed of the tape so less tape is used 

to record a given interval.

Continuous auto-reverse changes the 

tape head direction after one side of the tape has 

been used. This avoids the need for a researcher 

Figure 1. Wiring schematic for an automated frog call data logger. The components 

within the dotted box are required only for the voice time stamp.
89

Table 1. Primary components used in the sample automated frog call data logger

Items needed for each unit Model Approximate 

Price

Tape recorder SONY TCS-60DV Pressman 120

Microphone Shure Omnidirectional Dynamic 50

Recycle timer (Hour/minute) SSAC RS1A34 55

Recycle timer (Minute/second) SSAC RS1A12 55

12-volt battery 7Amp SLA 15

DC power converter Cigarette lighter adapter 10

Container 50-caliber ammunition can 5

Voice-time stamp Keychain voice clock 10

Timer/relay 20

Microphone1 0

Total $340

1The secondary microphone used for the voice-time stamp was included with the tape recorder.

Figure 2. Example of the interior layout of an automated frog call data logger; a.) analog tape recorder, 

b.) two solid state recycle timers, c.) 12v, 7amp sealed lead acid battery, d.) voltage regulator, e.) voice 

stamp assembly, f.) 1⁄4" microphone female jack, g.) 1⁄4" male microphone jack on microphone cable, 

h.) 50-caliber ammunition can.
90

to flip tapes over before the first side is 

spent. This feature varies from standard 

auto-reverse in that continuous autoreverse functions while the tape recorder 

is in record mode and standard operates 

only in play mode.

b. One or two 12-volt timer(s) are needed 

to run each automated frog call data logger. If the automated frog call data logger 

is intended to sample continuously (24-

hr/day), then a single (minute/second) 

timer is necessary. If a specific period 

within a day is desired a second (hour/

minute) timer is required. Solid-state 

encapsulated recycle timers have been 

widely used in automated frog call data 

loggers. These timers are programmed 

by adjusting two series of binary 

switches, one series for "ON" time and 

the other series for "OFF" time. The programmer must make absolutely sure the 

combined "ON" and "OFF" times equal 

1 hour for the minute/second timer and 

24 hours for the hour/minute timer. 

Greater detail regarding timer programming and technical data are available at 

the supplier's website (www.ssac.com). 

Other types of timers (555, BioQuip 12v 

DC timer) are available and have been 

used with success but require advanced 

knowledge of electronics, are less flexible to program, and can be less reliable.

c. Any single or combination of batteries

totaling 12 volts will suffice. The greater 

the amperage the longer the automated 

frog call data logger can operate without 

changing or replacing the batteries. 

Rechargeable batteries are recommended 

including the 12-volt, 7-amp sealed lead 

acid battery (SLA) illustrated in the sample automated frog call data logger 

(fig. 2). If multiple batteries are used, a 

battery holder is recommended. No 

batteries are used in the tape recorder as 

the main battery powers the entire unit.

d. Although the recycle timers run on 12v, 

tape recorders typically require 3 to 6v 

DC power. Rather than build a voltage 

step-down, this design uses an automotive cigarette lighter adapter capable of 

converting from 12v to 9, 7.5, 6, 4.5, and 

3v. The DC power input of the cigarette 

lighter adapter can be modified by cutting 

off the cylinder and contacts and splicing 

the timer outputs directly to the adapter 

input leads. The power adapter output 

will be connected to the external power 

jack or directly to the battery connections 

of the tape recorder. Attaching the power 

directly to the tape recorder battery terminals provides a more reliable unit than 

using the external power jack due to the 

small surface area of the external jack. 

This connection can easily be made by 

building insulated dummy batteries with 

the power connections at the ends (fig. 3).

e. The main purpose of a container is to 

protect the electronics from the elements 

in any easy-to-transport package. 

Figure 3 shows a very economical (< $5) 

surplus 50-caliber ammunition can. 

Alternatives include plastic pails, toolboxes, tackle boxes, and Otter® or Pelican® cases.

f. Microphones should be omni-directional and should not require an additional power source. In most cases 

monaural models are the only choices 

given these criteria, but they work adequately. Superior recordings are possible 

with DC powered stereo microphones, 

although the relatively short battery life 

can increase the maintenance schedule of 

the data loggers. The microphone cable 

can be passed through a port placed in the 

container or a microphone jack can be 

installed in the side of the container. 

Microphones can be shielded from the 

elements by placing them inside a cut-off 

plastic soda bottle. Additionally, foam 

can be placed between the bottle and the 

microphone head to reduce wind noise 

but this can introduce moisture wicking.

g. Most tape recorder manufacturers recommend tapes no longer than 90-minutes, 

because tapes of greater length are too 

thin and stretch under the tension of 
91

recording, thus reducing recording quality and reliability.

h. An optional voice-time stamp is highly 

recommended especially if calling phenology is of interest. This feature allows 

the researcher to know the time of each 

recorded interval with reasonable precision and makes reviewing the tapes considerably more easy. A voice-time stamp 

is made by triggering a talking clock at 

the same time the tape recorder is activated. This is accomplished by splitting 

the timer output to both the tape recorder 

and a relay that triggers the voice clock. 

To prevent the voice clock from continuously announcing the time during a 

recording, the relay must be supplied 

power only briefly at the beginning of the 

recording period. This can be accomplished by building a binary logic circuit 

or by using a timer. A second microphone picks up the announcement of the 

voice clock while the tape recorder 

records signals from both the primary 

external microphone and this smaller secondary internal microphone. The internal 

microphone should be placed away from 

the tape recorder and near the voice clock. 

In the example data logger, the voice 

clock, secondary microphone, and relay 

are housed in their own container and are 

Figure 3. Example of a dummy battery, an external power jack alternative. The battery 

door on this tape recorder has been notched to allow clearance for the wires. Wooden 

dowels (3/8", 0.9525cm) were used as insulators between the battery terminals.

at the opposite end of the can from the 

tape recorder (figs. 2-4).

2. Tools

a. Wire cutters for cutting rolls of wire 

into shorter lengths.

b. Wire strippers for removing the insulation from the ends of the wires.

c. Wire crimps are necessary for making 

wireless connections.

d. A drill or Dremel® can be useful for 

making modifications to the container.

e. A soldering iron is necessary for making solder connections and should be 

used to prepare multistrand wire for 

solderless connections.

f. A multimeter can be very helpful in 

troubleshooting connections as well 

as checking and maintaining 

batteries.

g. A 12v automotive battery charger can 

be used to charge a single battery or 

to run a bank-charging system 

(fig. 5). If a bank-charger is used, 

each battery should be individually 

fused and the fuse rating should be 

less than the maximum amperage of 

the battery and greater than the 

charge amperage. This will allow the 

batteries to be charged without blowing fuses. If a battery does short, 

however, it will blow only its fuse.
92

Figure 4. Example of a voice stamp assembly. The voice clock (far right) and timer/

relay are secured in a plastic food container and the microphone is mounted to the lid. 

Enclosing the assembly in its own container helps isolate the microphone from the 

sound of the tape recorder running.

3. Consumable materials

a. Stranded hook-up wire (18AWG). 

Multiple colors can be helpful to prevent confusion in polarity.

b. Female terminal connectors (1/4")

c. Light-duty 60/40 rosin core solder

(0.050" diameter).

d. Heat-shrink tubing or liquid tape.

4. Assembly

a. Charge all the batteries if using 

rechargeable batteries.

b. Make any modifications to the container that may be necessary (for 

example, drilling holes for ports or 

jacks) and install the appropriate 

hardware.

c. Program all timers to the desired schedule.

d. Dry fit all the individual components in 

the container to determine the best 

placement and layout.

e. Build the wiring loom to accommodate 

the location of the components.

f. Outside the container, attach the wiring 

loom to all the components except the 

battery.

g. Connect the battery to test the unit and 

make necessary corrections until the 

unit operates.

h. Disconnect the battery and transfer the 

partially assembled unit into the container.

i. The automated frog call data logger is 

now ready for use.

Setup

1. In the lab or office

The timers begin cycling when power is 

applied; therefore, make the power 

connections at the time the automated frog 

call data loggers are intended to begin recording. The voice clock should announce the 

correct time and the LED on the voltage stepdown should remain on for the duration of the 
93

programmed recording interval. In the 

example automated frog call data logger the 

batteries would be turned on at 06:00. At this 

point the timers would be cycling but the unit 

will not record until the record button is 

depressed in the field. This step should be 

performed the day before field deployment. 

Make sure the tape recorder is turned off. 

Label and insert a cassette tape into the 

recorder. Make sure the tape is rewound, on 

side "A," and the tape recorder, if it has autoreverse, is set in the correct direction. If there 

is any interruption of power to the timers they 

will reset to the time the power was reapplied.

2. In the field

Depress the record button on the tape 

recorder, connect the microphone, and close 

Figure 5. Example of a six battery bank charger. Each battery is individually fused to prevent 

catastrophic failure should an accidental short occur.

the case. The unit will not begin to record 

until the time the timers were started the preceding day. The microphone should be 

secured to woody vegetation or to a microphone stand if there is no structure available. 

It is important to place the microphone near 

the breeding site but the main unit can be 

placed anywhere the microphone cord can 

reach, which should be a secure site, away 

from possible flooding or vandalism. The 

data loggers can be locked closed and 

secured a tree or other sturdy object. Notify 

managers as to the location and appearance 

of the data loggers, because they could be 

easily mistaken as an explosive device 

(fig. 6). Tapes can be changed in the field, 

but it is not practical to change batteries 

without retrieving all the units.
94

3. Listening to tapes

Tapes can be reviewed at any time, and 

depending on the number of units deployed, it is 

easy to accumulate a backlog of tapes. Tape 

review should be conducted in a quiet area with 

as few distractions as possible. Listening 

requires about twice the recording time. All 

observations should be recorded on a data sheet 

(see fig. 7 for a sample data sheet). Important 

data fields should include the site, recording 

dates, time of each interval, and species calling. 

Additional data could include NAAMP call 

rank (fig. 7). Observations of sounds other than 

frog calls (for example, rain falling on the 

microphone, aircraft) can be useful in the interpretation of the tapes and should be noted.

Figure 6. Automated frog call data logger setup near a small pond.

Tips

• Twist and tin (apply small amount of solder) 

all stranded wires before crimping if using 

solderless connectors, and use heat-shrink 

tubing to cover and protect all solder and 

crimp joints. This greatly improves the 

reliability, durability and longevity of the 

data logger.

• Start by building a single prototype unit and 

after all the "bugs" have been worked out, 

use an assembly line technique to speed the 

process of building the others.

• Consider the research questions/objectives 

before programming and deploying your 

data loggers. Listening to the tapes can be 

very time consuming, so recording time 

should be minimized 

while still meeting 

research goals.

• Use high quality headphones that completely 

cover the listeners' ears 

for reviewing the 

recorded material. 

While doing so, try not 

to multitask, because it 

can be easy to overlook 

a call if the listener is 

distracted.

• If recordings were made 

using an extended 

record time feature, 

then playback must be 

performed on a like 

unit. This could 

require purchasing 

additional tape recorders to review the tapes.
95

Figure 7. Sample data sheet used to review automated frog call data logger tapes from Okefenokee National 

Wildlife Refuge.

Okefenokee NWR Frog Logger Data Sheet

Site Name/Number: __________________ Dates: ____________________

Logger Number: __________________ Listener 1: ___________________

Listener 2: ___________________

Int/Hour Night 1 Night 2 Night 3 Night 4

1/

2/

3/

4/

5/

6/

7/

8/

9/

10/

11/

12/

13/

14/

** include species heard and NAAMP calling codes at each interval in parentheses

0= no frogs can be heard calling; 1= individual calls not overlapping; 2= calls are overlapping; but 

individuals are still distinguishable; 3= numerous frogs can be heard; chorus is constant and 

overlapping

Notes:
96

SELECTED REFERENCES

Bridges, A.S., and Dorcas, M.E., 2000, Temporal 

variation in anuran calling behavior: Implications 

for surveys and monitoring programs: Copeia 

2000, p. 587-592.

Mazanti, L.E.,1999, The effects of atrazine, metolachlor 

and chlorpyrifos on the growth and survival of 

larval frogs under laboratory and field conditions: 

Unpublished Ph.D. dissertation, University of 

Maryland.

Peterson, C.R. and Dorcas, M.E., 1992, The use of 

automated data-acquisition techniques in monitoring amphibian and reptile populations, in

McCullough, D.R., and Barrett, V., eds., Wildlife 

2001: Populations: London, Elsevier, Applied 

Science, p. 369-378.

Peterson, C.R., and Dorcas, M.E., 1994, Automated 

data acquisition, in Heyer, W.R., Donnelly, M.A., 

McDiarmid, R.W., Hayek, L.C., and Foster, M.S., 

eds., Measuring and monitoring biological 

diversity: Standard methods for amphibians: 

Washington, DC, Smithsonian Institution Press, 

p. 47-57.

Rand, A.S., and Drewry, G.E., 1994, Acoustic monitoring at fixed sites, in Heyer, W.R., Donnelly, M.A., 

McDiarmid, R.W., Hayek, L.C., and Foster, M.S., 

eds., Measuring and monitoring biological 

diversity: Standard methods for amphibians, 

Washington, DC, Smithsonian Institution Press, 

p. 151-152.

RELATED WORLD WIDE WEB SOURCES

http://www.uga.edu/srelherp/ecoview/Eco19.htm

http://www.parcplace.org/education/techniques/froglogger.htm

http://www.bedfordtechnical.com/index.htm

http://www.bio.davidson.edu/people/midorcas/research/StResearch/Cocklinetal00/Cocklinetal.htm

http://www.bio.davidson.edu/people/midorcas/research/StResearch/Briggs99/scottprop.html

http://www.bio.davidson.edu/people/midorcas/research/herppub-pres/dorcas-pdfs/MohrDorcas1999.pdf

http://www.bio.davidson.edu/people/midorcas/research/herppub-pres/dorcas-pdfs/PetersonDorcas1994.pdf

http://www.bio.davidson.edu/people/midorcas/research/herppub-pres/dorcas-pdfs/PetersonDorcas1992.pdf

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PLATES
98
99

Amphibians
100

Marbled Salamander (Ambystoma opacum). 

Stupka’s Sinkhole Pond, Cades Cove.

Spring Salamander (Gyrinophilus 

porphyriticus). Andrews Bald.

PLATE 1

A

B
101

Shovel-nosed Salamander (Desmognathus 

marmoratus). Mill Creek, Cades Cove.

Jordan=s Salamander 

(Plethodon jordani). 

Clingmans Dome.

PLATE 2

A

B
102

American Toad (Bufo 

americanus). Cades Cove.

Wood Frog (Rana sylvatica). 

Gregorys Cave, Cades Cove.

PLATE 3

B

A
103

Habitats
104

B

View southwest from Clingmans Dome. 

Fontana Reservoir is in the distance.

Mt. LeConte.

PLATE 4

A
105

Top of Andrews Bald, view toward 

Thunderhead Mountain.

Clingmans Dome (highest point in the Great 

Smokies). The effects of clearcut logging and 

fire are clearly visible on the right; the dead 

trees are Fraser Fir, killed by the Balsam woolly 

adelgid.

PLATE 5

A

B
106

Cades Cove, view to southeast.

Cades Cove, view toward Gregorys Bald (hidden in clouds). Friendly resident.

PLATE 6

A

B
107

Larvae and Tadpole Mouthparts
108

Spotted Salamander (Ambystoma maculatum)

Marbled Salamander (Ambystoma opacum)

Mole Salamander (Ambystoma talpoideum)

PLATE 7

A

B

C
109

Hellbender (Cryptobranchus alleganiensis)

Shovel-nosed Salamander (Desmognathus marmoratus)

Seal Salamander (Desmognathus monticola)

Imitator Salamander (Desmognathus imitator)

PLATE 8

Spotted Dusky Salamander (Desmognathus conanti)

A

B

C

D

E
110

PLATE 9

Black-bellied Salamander (Desmognathus quadramaculatus)

Ocoee Salamander (Desmognathus ocoee)

Junaluska Salamander (Eurycea junaluska)

Three-lined Salamander (Eurycea guttolineata)

Santeetlah Salamander (Desmognathus santeetlah)

A

B

C

D

E
111

Cave Salamander (Eurycea lucifuga)

Long-tailed Salamander (Eurycea longicauda)

Four-toed Salamander (Hemidactylium scutatum)

Spring Salamander (Gyrinophilus porphyriticus)

Blue Ridge Two-lined Salamander (Eurycea wilderae)

PLATE 10

A

B

C

D

E
112

Common Mudpuppy (Necturus maculosus)

PLATE 11

Eastern Red-spotted Newt (Notophthalmus viridescens)

Mud Salamander (Pseudotriton montanus)

Black-chinned Red Salamander (Pseudotriton ruber)

A

B

C

D
113

Northern Cricket Frog (Acris crepitans)

American Toad (Bufo americanus)

Fowler’s Toad (Bufo fowleri)

PLATE 12

A

B

C
114

Eastern Narrow-mouthed Toad 

(Gastrophryne carolinensis)

Cope’s Gray Treefrog (Hyla chrysoscelis)

Spring Peeper (Pseudacris crucifer)

PLATE 13

A

B

C
115

Upland Chorus Frog (Pseudacris feriarum)

American Bullfrog (Rana catesbeiana)

PLATE 14

A

B
116

Northern Green Frog (Rana clamitans)

Pickerel Frog (Rana palustris)

PLATE 15

B

A
117

Northern Leopard Frog 

(Rana pipiens)

Eastern Spadefoot (Scaphiopus holbrooki)

Wood Frog 

(Rana sylvatica)

PLATE 16

A

B

C
118
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