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Department of 5 4 5 Agriculture **. 

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Pacific Northwest 

Research Station 

Skmpling Methods .fo.r 

Terrestrial A.mphibians 

General Technical 

Report 

PNW-GTR-256 

July 1990 / 

and R.eptiles 

Paul Stephen Corn and R. Bruce Bury 
Wildlife-Habitat Relationships:

Sampling Procedures for

Pacific Northwest Vertebrates

Andrew B. Carey and Leonard F. Ruggiero, Technical Editors

Sampling Methods for Terrestrial

Amphibians and Reptiles

Paul Stephen Corn

Zoologist

U.S. Department of the Interior

Fish and Wildlife Service

National Ecology Research Center

4512 McMurray Avenue

Fort Collins, Colorado 80525-3400

R. Bruce Bury

Research Zoologist

U.S. Department of the Interior

Fish and Wildlife Service

National Ecology Research Center

4512 McMurray Avenue

Fort Collins, Colorado 80525-3400

USDA Forest Service

Pacific Northwest Research Station

Portland, Oregon

General ‘Technical Report PNW-GTR-256

1990
Preface Concern about the value of old-growth Douglas-fir forests to wildlife in the Pacific

Northwest began escalating in the late 1970s. The available information on wildlifehabitat relationships suggested that as many as 75 species including amphibians,

birds, and mammals, could be dependent on old-growth forests. The USDA Forest

Service chartered the Old-Growth Forest Wildlife Habitat Program to investigate the

role old growth plays in maintaining viable populations of wildlife. It was apparent that

broad surveys of vertebrate communities would be necessary to determine which

species were truly closely associated with old-growth forests. Insufficient guidance on

techniques, procedures, and sample sizes was available in the existing literature. We

assembled a team of researchers from universities and Federal agencies to conduct

pilot studies to develop sampling protocols and to test the basic experimental design

for contrasting the wildlife values of young, mature, and old-growth forests. The

sampling protocols resulting from the pilot studies were implemented in 1984-86

across broad areas of the Cascade Range in southwestern Washington and in

Oregon, the Oregon Coast Ranges, and the Klamath Mountains of southwestern

Oregon and northern California. Naturally, improvements were made to the protocols

as time passed. A tremendous amount of experience in sampling was gained.

Our goal in this series is to compile the extensive experience of our collaborators into

a collection of methodology papers providing biologists with pilot study-type information for planning research or monitoring populations. The series will include papers

on sampling bats, aquatic amphibians, terrestrial amphibians, forest-floor mammals,

small forest birds, and arboreal rodents, as well as papers on using telemetry for

spotted owl studies and a guide to bird calls.

Andrew B. Carey

Leonard F. Ruggiero
Abstract Corn, Paul Stephen; Bury, R. Bruce. 1990. Sampling methods for terrestrial

amphibians and reptiles, Gen. Tech. Rep.. PNW-GTR-256. Portland, OR: U.S.

Department of Agriculture, Forest Service, Pacific Northwest Research Station.

34 p.

Methods described for sampling amphibians and reptiles in Douglas-fir forests in

the Pacific Northwest include pitfall trapping, time-constrained collecting, and surveys

of coarse woody debris. The herpetofauna of this region differ in breeding and nonbreeding habitats and vagility, so that no single technique is sufficient for a community study. A combination of pitfall trapping and hand collecting is the most

effective approach.

Keywords: Amphibians, reptiles, sampling techniques, pitfall trapping, timeconstrained collecting, downed wood.
Contents 1 Introduction

3 Objectives

3 Overview

5 Time-Constrained Searches

6 Surveys of Coarse Woody Debris

6 Pitfall Trapping

7 Experimental Design

7 Time-Constrained Searches

7 Surveys of Coarse Woody Debris

8 Pitfall Trapping

11 Field Methods

11 Crew Sizes

11 Time Frame and Weather

12 Operating Guidelines

17 Identification

17 Disposition of Specimens

18 Data Analysis

21 Conclusions

21 Acknowledgments

22 Equivalents

22 Literature Cited

27 Appendix 1

28 Appendix 2

28 Data Sheets

32 Appendix 3

32 Materials Needed for TCS or Surveys of CWD

32 Materials Needed for Pitfall Installation and Operation

34 Materials Needed in the Lab
Introduction There is a rich herpetofauna in the Pacific Northwest, with 43 species of amphibians

and reptiles present west of the Cascade Range (appendix 1). Depending on the geographical area, 19 to 32 species may be present at a given site (fig. 1). The number

of species of amphibians is consistent at 13 to 15 species in most areas in this region, but reptiles range from 5 species in southwestern Washington to 17 species in

both southwestern Oregon and northwestern California. The difference is due to increased aridity and higher temperatures in the southern locales, which favor reptiles.

Although a diverse reptilian fauna may occur in an area, many species (particularly

snakes) are locally rare or restricted to certain habitats; for example, oak-woodland

(many snakes) or permanent water (turtles).

FS LSn7

\

i

\

Figure l-Number of amphibians and reptiles potentially present

in different regions of the Pacific Northwest west of the crest

of the Cascade Range. The histograms are by major taxonomic

groups: F = frogs, S = salamanders, L = lizards, Sn = snakes,

and T = turtles.
During recent research in western Oregon and Washington, we found few or no

reptiles present in closed-canopy Douglas-fir (Pseudotsuga menziesii (Mirb.) France)

forests (Bury and Corn 1987, 1988; Corn and Bury, in press). Reptiles are usually

encountered in rocky, open areas (for example, cliff faces) or in grasslands and oak

woodlands (Herrington 1988, Nussbaum and others 1983); these habitats were rare

or absent in the forest stands we studied. Thus, reptiles were a small fraction of the

sampled herpetofauna, and they will receive little mention here. Biologists will need

to employ special techniques if reptiles are encountered at a study site (see Bury and

Raphael 1983, Jones 1986, Scott 1982b).

In the Pacific Northwest, amphibians are often found in terrestrial habitats, particularly in forests, and among terrestrial vertebrates may be the most numerous group. Terrestrial salamanders, for example, can exceed over five individuals/m* in local aggregations (Bury and Raphael 1983, Jaeger 1979). In 1983, Bury estimated that there

were over 400 salamanders/ha in old-growth redwood forests in northern California

(Bury 1983). In 1984, Raphael reported densities of 10 to 180 salamanders/ha in

Douglas-fir forests in northern California (Raphael 1984). We estimated that mean

density of plethodontid salamanders associated with downed wood ranged from

364/ha in young Douglas-fir forests to 744/ha in old-growth forests (Corn and Bury, in

press). For eastern deciduous forests in New Hampshire, Burton and Likens (1975)

estimated about 3,000 salamanders/ha, and Hairston (1987) estimated that energy

present in salamanders in southern Appalachian forests exceeds that of all other

vertebrate predators combined.

Amphibians are important components of the northwestern fauna in ways other than

numbers or biomass. Of 22 amphibian species inhabiting forest habitats in the Pacific

Northwest, 14 species (64 percent) are endemic (species whose distributions are restricted to the Pacific Northwest). Many of these habitats are affected increasingly by

human activities.

Several species of plethodontid salamanders are more abundant in older forests, or

show relations to habitat features that are prominent in old-growth forests. Ensatinas1

are more abundant in older Douglas-fir forests than in younger stands in northern

California (Raphael 1984). Ensatinas, Oregon slender salamanders, and clouded salamanders are often associated with large pieces of downed wood (Aubry and others

1988; Bury and Corn 1988; Corn and Bury, in press). Coarse woody debris (CWD) is

a major component of old-growth forests and is severely reduced by modern forestry

practices (Harmon and others 1986, Maser and Trappe 1984). The plethodontid salamanders in general are useful for assessing logging impacts because they have completely terrestrial life cycles (the eggs are deposited on land and hatch into miniature

individuals), and most species have stable populations (Hairston 1987).

1

 Scientific names of reptiles and amphibians are given in

table 6 (appendix 1).
Objectives

Overview

The relations of frogs and aquatic-breeding salamanders to older forests are more

difficult to explain than are the relations of plethodontids. Most of these species use

terrestrial habitats to a degree, especially for feeding. They also may migrate overland to breeding ponds or streams and, thus, temporarily occur in many habitats

during their travels. Tailed frogs previously had been considered to be closely tied to

streams (Metter 1967) but we discovered that they are found in forests long distances from flowing water (Bury 1988). Our results also suggest that juvenile tailed

frogs disperse into terrestrial habitats away from streams.

Given the diversity of amphibian life histories, habitat preferences, and different

means of locomotion, more than one sampling technique is needed to sample adequately all species of amphibians. We used several methods to sample amphibians;

methods for sampling aquatic species are discussed separately (Bury and Corn, in

press). We sampled the terrestrial herpetofauna in three main ways: (1) time-constrained searches (TCS), (2) searching specified numbers of pieces of downed wood

(CWD surveys), and (3) pitfall trapping.

We will discuss the objectives, sampling design, and techniques specific to each

method separately. We will then discuss techniques common to all the methods we

used and make recommendations for effectively sampling the herpetofauna in the

Pacific Northwest.’ The methods described here were used by the Old-Growth Forest

Wildlife Habitat Program (Ruggiero and Carey 1984) in field work from 1983 to 1985.

This program included studies of vertebrates in Douglas-fir forests in California,

Oregon, and Washington west of the Cascade Range (Ruggiero and others, in press).

With the exception of experiments to determine the most effective design for pitfall

trapping (Bury and Corn 1987), these methods were not rigorously tested against

alternatives (field methods, particularly hand-collecting techniques, have rarely been

subjected to experimental comparisons). Rather, they reflect our current professional

judgment and draw heavily from other recent descriptions of field methods (Campbell

and Christman 1982, Jones 1986, Raphael and Barrett 1981, Vogt and Hine 1982).

The primary objective of our study was to identify species associated with old-growth

Douglas-fir forests (Ruggiero and Carey 1984), and so the techniques we used were

slanted to favor survey methods. Pitfall trapping and CWD surveys will provide some

information on populations. These data can be used to analyze habitat use by individual species and the patterns shown by groups of species in different habitats. Coarse

woody debris surveys and TCS can also provide detailed information on the use of.

microhabitats by various species. Basic ecological data are needed that can be applied to recommendations for management of specific habitats.

There are marked differences in catch between hand collecting (TCS and CWD ‘-

surveys) and pitfall trapping (table 1). Species such as clouded salamanders and

Oregon slender salamanders are closely associated with CWD and were frequently

caught by hand but were trapped infrequently. Tailed frogs, newts and other migratory

species were trapped effectively in pitfalls but rarely were caught by hand.

The choice of a specific method to achieve stated objectives depends on the species

under study as well as the scope of the objectives. If a small-scale study on one or a

few species is intended, then only one method may be needed. A survey of community structure over a large geographic area will likely require all three methods.

3
Table 1-Comparison of captures of amphibians and reptiles by pitfall trapping

and time-constrained searches (TCS), H.J. Andrews Experimental Forest, 1983

Number of captures

Pitfallsb

Species TCSa

Total 281

Summerc

206

Fallc

822

TCS

Percent of captures (rank)

Pitfalls

Summer Fall

Salamanders:

Northwestern salamander

Pacific giant salamander

Clouded salamander

Oregon slender salamander

Ensatina

Dunn’s salamander

Rough-skinned newt

0 (-) 0 (-) 5 (5)

0 (-) 1 (13) 2 (6)

28 (2) 3 (10) 1 (8)

22 (3) 4 (7) 1 (10)

43 (1) 24 (1) 25 (2)

1 (5) 1 (14) 1 (9)

1 (6) 15 (2) 37 (1)

Frogs:

Tailed frog

Pacific treefrog

Red-legged frog

0 (-) 9 (5) 19 (3)

2 (4) 4 (8) 1 (7)

0 (-) 1 (12) 6 (4)

Lizards:

Western skink

Northern alligator lizard

Western fence lizard

1 (8) 11 (4) 1 (12)

1 (9) 13 (3) 1 (11)

1 (7) 3 (11) 1 (15)

Snakes:

Rubber boa

Northwestern garter snake

Common garter snake

1 (15) 0 (-)

7 (6) 1 (14)

4 (9) 1 (13)

a

 TCS were done for 8 staff hours in 18 study areas in April.

b

 Arrays of pitfall traps with drift fences (Bury and Corn 1987) were operated in the same areas for 180

days from late May to November.

c

 The results of pitfall trapping are divided into the first 90 days of trapping (summer) and the second 90

days (fall).

Throughout this paper, we will use the terms study site and stand interchangably.

This is due to the bias of working in forests, where study sites tend to encompass

areas of more or less uniform habitat, which are referred to as stands. Stands in the

old-growth studies were patches of forest of uniform age with a minimum area of

10 ha (Carey and Spies, in press).

-
Time-Constrained

Searches

Time-constrained searches involve searching study areas for amphibians and reptiles, which are immediately collected by hand (Bury and Raphael 1983, Campbell

and Christman 1982). Equal effort is expended in each area searched, as measured

by the number of staff hours spent searching. Thus, each search will have a specific

time limit, dependent on the prescribed effort and the crew size. Time-constrained

searches are most useful for determining presence or absence of species and for

providing initial data on the types of microhabitats occupied by individual species.

Time-constrained searches are not suitable for providing population data beyond presence or absence. Because this is a “plotless” technique, the same amount of potential

habitat tends to be searched in each study area: however, amounts of suitable habitat differ among study areas. Results from some TCS may show habitat-poor areas

yielding similar numbers of animals as habitat-rich areas, even though the population

sizes may be quite different. Indeed, evidence is that salamanders are more clumped

in areas with less habitat, which will increase the bias in favor of these areas. In the

Coast Ranges of Oregon, we found the density in downed wood (number per m

3

) of

ensatinas was significantly higher in young and mature stands compared to old

growth (fig. 2) (Corn and Bury, in press). In this case, TCS could possibly result in an

inverse relation of numbers caught to’ actual population size.

If population estimates are an objective, then other techniques need to be applied.

We used CWD surveys effectively (explained below), but another common method is

complete removal of all residents of a predetermined area (Bury 1983; Campbell and

Christman 1982; Jaeger 1979; Raphael 1984; Scott 1976, 1982a). Plot searches are

labor intensive: Bury (1983) required 20 to 44 staff hours to search 0.125-ha plots in

old-growth redwood forests in northern California. For surveys of several study areas,

plot searches may require too much effort to produce sample sizes large enough for

statistical analysis.

6 -

5 -

4 -

l

VOLUME OF DOWNED WOOD (CUBlC METERS/HECTARE)

Figure 2-Salamander density in

downed wood. The density of

salamanders (number/m3

) in

downed logs is inversely related

to the amount of downed wood

present in the study areas.

Salamanders appear to be less

clumped as more habitat is available.

5
For initial surveys of presence or absence, TCS are more effective than plot searches

because collectors are free to examine large objects over a wide area, and usually

more amphibians are found in large objects than in the leaf litter, at least in the

Pacific Northwest. This method is efficient because the objects searched are most

likely to yield animals. In northwestern forests, TCS may produce as much as a 10

times greater yield than will area-constrained collecting (Bury and Raphael 1983). In

recent studies, capture rates of TCS have ranged from one to two animals per staff

hour in the Cascade Range in Oregon and Washington (Aubry and others 1988, Bury

and Corn 1988), to over eight animals per staff hour in northern California (Welsh

and Lind 1988). Time-constrained searches are best employed when several study

areas need to be surveyed in a short time.

Surveys of Coarse

Woody Debris

In 1985, we were confronted with the choice of initiating TCS in the Oregon Coast

Ranges or developing a technique to quantify habitat use and estimate density of

selected species of salamanders. We chose the latter and developed a technique

involving searches of predetermined numbers of pieces of downed wood. Numbers of

animals caught were then related to the amounts of CWD in the stand, and minimumdensity estimates were calculated.

Surveys of CWD are operationally similar to TCS; but to estimate animal densities,

the density of CWD must be known. Knowing the amount of CWD present also allows

for quantifying microhabitat use and drawing meaningful comparisons of microhabitat

use among species.

The primary drawback of surveys of CWD is that density estimates apply to only

one feature of the habitat. Surveys of CWD underestimate density of species using

downed wood only occasionally; for example, most species of woodland salamanders

(Plethodon spp.) frequent rocky soils, but an unknown fraction of a population may

occur in CWD. For species strongly associated with CWD (for example, the clouded

salamander or the Oregon slender salamander), surveys of CWD should provide

general estimates of population sizes.

Pitfall Trapping Pitfall trapping is a flexible technique that can be used to achieve several objectives;

for example, drift fences with pitfall traps have been used to encircle specialized habitats such as amphibian breeding ponds (Gibbons and Semlitsch 1981, Shoop 1968,

Storm and Pimentel 1954). This technique can be used for complete enumeration of

breeding populations. Pitfall trapping also has been employed widely for surveys of

amphibian and reptile diversity and abundance in different habitat types (Bury and

Corn 1987; Campbell and Christman 1982; Friend 1984; Jones 1981, 1986; Raphael

1984; Vogt and Hine 1982; also see selected papers in Ruggiero and others, in press;

and Szaro and others 1988). The main drawback of pitfall trapping is that trapability

differs widely among species (Bury and Corn 1987, Campbell and Christman 1982,

Gibbons and Semlitsch 1981). A survey of ail species of herpetofauna in an area

therefore requires more than one technique.
Pitfall trapping provides data on the presence or absence of species, and because

the trapping effort can be quantified and standardized across study areas, relative

abundances can be calculated. Estimates of actual population size may be possible,

though probably only for abundant species. Pitfalls may be used as live traps if

checked frequently, and mark and recapture techniques also may be used. If pitfalls

are used as a removal method to estimate density, then the area being trapped must

be known. This is extremely difficult to determine for most herpetofauna and is

something we have not done in any of our studies.

Pitfall trapping is also useful for investigating seasonal activity patterns. Traps can be

operated continuously, so that variation in activity due to weather can be detected

(Bury and Corn 1987). Pitfall traps are permanent structures, so long-term monitoring

can be accomplished by operating the same trap array or grid periodically over

several years. Trapping has unknown effects, however, on population structure due

to the removal of resident individuals.

Experimental

Design

This technique is a quick survey method requiring few restrictions on the approach.

Three points need to be considered: (1) collecting should be done away from forest

Time-Constrained

Searches

edges; (2) aquatic habitats, such as breeding ponds or creeks should be avoidedthese are covered by a separate protocol (Bury and Corn, in press); and (3) collecting

should cover as much of the stand as possible. There are two ways to accomplish

this last point. One is to devote enough time to the search to be able to collect across

the entire study area. The second is to restrict the search to a fairly small area (for

example, a circle with a radius of 25 m) and restrict the amount of time spent collecting. The number of smaller areas that can be searched in each study area depends

on the amount of time devoted to the TCS. We found that 6 or 8 staff hours of collecting were sufficient; few additional species were detected by collecting for longer than

that. If 1 hour is spent in each of the subsamples, then six to eight areas can be

searched in each study area.

Surveys of Coarse

Woody Debris

This technique is somewhat more complicated than TCS in that it involves systematicaily searching a predetermined number of logs in each study area. Several questions must be addressed when a study is designed, including how many logs to

sample, how to apportion the sample among the different decay states of downed

wood, and how to select the logs sampled.

in 1985 in the Oregon Coast Ranges, we conducted CWD surveys in 18 study areas.

Each survey included 30 logs greater than 10 cm in diameter. We selected 10 logs

in decay classes 1 and 2, 10 in decay class 3, and 10 in decay classes 4 and 5 (see

Franklin and others [1981] or Maser and Trappe [1985] for methods of classifying

CWD). The three decay categories that we used reflected natural divisions of the

five-class scale. Class 1 and 2 logs are intact with more or less complete bark cover.

Class 3 logs have decaying sapwood, and the bark is beginning to slough off. Class

4 and 5 logs are thoroughly decayed, have little bark, and are disintegrating. We

recommend sampling equal numbers of logs in each of these decay categories. We

searched each log for a maximum of 20 staff minutes.

7
Pitfall Trapping 

We found salamanders in only 37 percent of the logs (198 of 536) that we examined, 

so a sample size of 30 logs per stand should probably be considered the minimum. if 

few logs are sampled and salamanders occupy a small percentage of these, then the 

estimates of salamander density will be based on minimal information. 

Logs to be sampled are best selected by a systematic sampling scheme (Mendenhaii 

and others 1971). If the study area has not been mapped and the locations of ail logs 

determined, it will not be possible to draw a random sample. A systematic sample 

involves selecting logs in a specified order as they are encountered while the crew 

moves through the stand. 

Planning pitfall trapping mainly involves selecting the appropriate trap design. We 

used two different pitfall designs in our old-growth studies (fig. 3). in 1983, we used 

arrays of pitfall traps with aluminum drift fences (Bury and Corn 1987). in 1984 and 

1985, we used grids of single pitfall traps without fences. There were quantitative and 

qualitative differences in the yield of each technique that must be considered in planning a project. 

PITFALL ARRAY 

DETAIL OF ONE ARM: 

Fr 

t 

Sm I 3m -I 

6 X 6 GRID OF PITFALL TRAPS 

COLUMN 

ABCOEF 

lx x xx x x 

2x x x x x x 

ROW 3 x x x x x x 

4x x xxx x 

5x x xx x x I 15m 

6x x XX x x 

llSm 

Place traps within 2 m of flagged station: 

use natural fenws (logs) where available 

Figure 3-Designs for arrangements of pitfall traps either with or without 

drift fences. PT = pitfall trap, FT = funnel trap. 
Trap rates for salamanders are similar for both arrays and grids, but arrays caught 

considerably more frogs and reptiles than grids did (table 2). The differences were 

due, in part, to the absence of drift fences in pitfall grids and the season when trapping was done. Pitfall arrays were open continuously for 180 days in 1983, from May 

to November. Grids were open for 30-50 days, beginning in October 1984. The grids 

were operated too late in the year to capture reptiles and large numbers of postmetamorphic juvenile frogs, which disperse from breeding sites in late summer or early 

fall. These frogs composed the majority of frogs caught by the arrays. 

Although arrays catch more animals than grids do, arrays are not necessarily better 

for determining presence or absence of amphibians. Grids caught few reptiles but 

were able to detect amphibians, including frogs, as well as or better than arrays 

(table 3). 

Table 2-Capture totals and trap rates (captures/100 trap nights) for major 

groups of amphibians and reptiles for 30 pitfall arrays in the Cascade Range of 

Oregon and Washington, 1983 (180 days), and for 48 pitfall grids in the Oregon 

Coast Ranges, 1984 and 1985 (80 days) 

Pitfall arrays Pitfall grids 

,. 

Group Species Captures Trap rate Species Captures Trap rate 

Salamanders 10 1145 1.77 8 1762 1.27 

Frogs 3 915 1.41 2 103 .07 

Lizards 3 79 .12 <.Ol 

Snakes 3 41 .06 0 

Table 3-Species of amphibians and reptiles inhabiting Douglas-fir forests at 

Old-Growth Wildlife Habitat Program study areas in Washington and Oregon 

Species 

Pitfall Pitfall 

array? grid? 

Central Southern Oregon 

Oregon Washington Coast 

Cascade Range Cascade Range Ranges 

Amphibians: 

Northwestern salamander 

Long-toed salamander 

Cope’s giant salamander 

Pacific giant salamander 

Olympic salamander 

Clouded salamander 

Oregon slender salamander 

Ensatina 

Dunn’s salamander 

Larch Mountain salamander 

Van Dyke’s salamander 

Western red-backed salamander 

Rough-skinned newt 

C 

P?b 

- 

P? 

C 

C 

P? 

P 

: 

- 

c 

c 

- 

C 

C 

C 

-*. 

c 

E 

c 

C 

- 

P? 

: 
Table 3-continued 

Species 

Pitfall Pitfall 

arraysa gridsa 

Central Southern Oregon 

Oregon Washington Coast 

Cascade Range Cascade Range Ranges 

Tailed frog 

Boreal toad 

Pacific tree frog 

Red-legged frog 

Yellow-legged frog 

Cascades frog 

Spotted frog 

Detection EfficiencyC 

Reptiles: 

Western skink 

Northern alligator lizard 

Southern alligator lizard 

Western fence lizard 

Rubber boa 

Sharp-tailed snake 

Ring-necked snake 

Gopher snake 

Sierra water snake 

Terrestrial garter snake 

Northwestern garter snake 

Common garter snake 

Western rattlesnake 

Detection Efficiency 

C 

rz 

C 

P? 

P? 

P 

59-77 

C 

cp 

C 

- 

P? 

P 

62-77 

C 

P 

: 

P? 

- 

- 

77-83 

C 

C 

c 

C 

- 

P 

- 

- 

P? 

C 

C 

- 

75-86 

c 

P? 

P 

P 

- 

- 

- 

- 

P? 

: 

- 

43-60 

F 

P 

P 

P 

P 

P? 

P 

P 

P 

P 

8 

b P= potential occurrence, C = captured in pitfall traps, - = species does not occur in the area. 

Potential occurrerm of a species in our study areas was uncertain. 

’ Species captured + species potentially present x 100. 

The choice of whether to install arrays or grids ultimately depends on the needs of 

the study. Arrays are superior for catching reptiles, but reptiles may not be abundant 

in forest habitats or of interest to the goals of a study. Arrays can provide large 

sample sizes in relatively shot-l periods. Grids remove fewer animals than arrays and 

may be more suitable for long-term monitoring. Both techniques are effective for 

catching small mammals as well as amphibians. 

Arrays may be placed in pairs, as we operated them in 1983 (fig. 3), or single arrays 

may be placed at more than one location within a stand. Three or four single arrays 

scattered throughout the stand may better assess the variation within study areas, but 

this approach requires significantly more time for checking the traps in each area. 

10 

L 
Field Methods

Crew Sizes

Time Frame and

Weather

Cost may be part of the decision on whether to install arrays or grids. Grids are not

substantially cheaper in cost of materials, because more pitfall traps can supplant the

cost of fencing. Grids took only about one-half the effort to install as the arrays did.

When personnel costs are high, this can result in a large difference in cost between

the two methods. The cost involved in checking the traps is similar and depends

mainly on the number of stands and the travel time between them.

This section provides instructions for carrying out TCS, surveys of CWD, and pitfall

trapping. We will not discuss selection of study areas. If the study is an integrated

wildlife survey, then study areas for mammals or birds can be used just as well for

studying the herpetofauna. All the techniques discussed here require small areas as

compared to bird or mammal studies.

Optimal crew sizes depend on the technique being used. Time-constrained searches

and surveys of CWD use the same collecting techniques, and three to four persons

are suitable for both. In both crews, one person is the data recorder, and the remaining people do the collecting. A 6-staff-hour TCS, done with a two-person crew plus a

recorder who does not collect, requires 3 hours, plus the time for breaks.

For pitfall trapping, a large crew is generally necessary to install traps, but only one

or two people are needed to check the traps once they are open. Installation of either

arrays or grids is relatively fast with a crew of six. Crews of this size can install two

arrays or grids per day. Two people can check a grid of 36 traps in an hour or less.

Several sites can be checked in one day, depending on the travel time between

study areas.

Hand collecting (TCS and surveys of CWD) should be done when amphibians are

most likely to be active; that is, in the rain. In the Pacific Northwest, this is either in

spring or fall (it rains in winter also, but low temperatures inhibit surface activity by

amphibians). If there are several study areas, then the primary consideration is that

the weather be as consistent as possible throughout the collecting period. Activity of

amphibians is highly dependent on weather, and comparisons between areas of collection under radically different weather conditions may not be valid. Collecting therefore should begin as early as possible in spring or as late as possible in fall, but still

avoiding lengthy periods of cold and snow. Collecting should not be done in heavy

snow; light snowfall in a period of wet weather probably will not seriously affect amphibian activity. Two TCS can usually be done in one day, but one survey of CWD

requires most of a day. It is possible, but not recommended, to split a survey between

two days.

Pitfall trapping has more flexibility, because all traps are open at the same time;

thereby reducing variability among study areas due to weather. The best season for

operating pitfalls depends on the animals being trapped. For amphibians, spring and

fall are again the periods of highest activity and will result in the largest catch. If

reptiles are being sampled, then early summer is the best time to open pitfall traps.

Pitfall installation can be done at any time, but data (Bury and Corn 1987) suggest

that pitfalls should be in the ground at least 1 month before trapping begins.

11
Operating Guidelines Time-constrained searches-Determine the number of 1-staff-hour searches that

can be done in the amount of time allotted to each study area. On a topographic map

or aerial photo of the study area, distribute the 1-hour searches for maximum coverage of the study area. The crew should enter each TCS with a map of the study area

that shows the approximate location of each l-hour search and the path to follow between searches, with compass headings and approximate distances. Each 1-hour

search should be confined to an area with a radius of about 25 m, and the center of

each 1-hour search should be at least 75 m from any forest edge.

Each TCS is a survey of as much habitat as possible within each study area. Move

from one object to the next after a few minutes. It is possible to spend over an hour

at one large log, but a maximum of 10 minutes per object should suffice. Assuming a

crew of two collectors and one recorder, each staff hour of search takes 30 minutes

of actual time. When an animal is found, time is spent by the collector in assisting

the recorder. The recorder should keep track of this time, and the total amount of

data recording time is added to the end of the search, so that 1 full hour of collecting

is achieved. This becomes more important in searches yielding many animals,

because data recording will require more time.

Surveys of CWD-The techniques involved here are more precise than those used

in TCS. Logs are chosen by a systematic sampling scheme. Specifically, a choice is

made to sample one log out of a certain number of logs encountered. In most habitats, choosing one out of every three logs will produce a survey covering a large

proportion of the study area. Further, logs are divided into subsamples based on the

decay state of the log. We compressed the standard five-point decay classification

into three categories: category A-decay classes 1 and 2, category B-decay class

3, and category C-decay classes 4 and 5. Sample 10 logs in each category (one of

every three logs encountered in each category) for a total of 30 in each study area.

Plot a path through the study area that will cover a large portion of the area but will

not intersect itself. For each decay category, choose a random number from one to

three. Begin following the designated path. At every downed log, determine the decay

category and whether the log should be sampled. The recorder keeps a running tally

of the number of logs encountered in each category. Each category of log accumulates at its own pace, and whether a log is sampled depends on the number of logs

encountered in that decay category. The decision may be, for example, to sample

every second category-A log, every third category-B log, and every first category-C

log. For this example, table 4 shows which logs will be selected from the first 20 logs

encountered.

When a log is selected, measure the total dimensions (see appendix 2 for data forms

and a description of the data to be recorded). Determine the tree species, if possible,

and the slope and aspect of the site where the log occurs. Search the log for a maximum of 20 staff minutes. Carefully remove any bark and tear into the decayed wood

layer by layer. If the entire log cannot be sampled within the time limit, search a portion of the log as completely as possible. This is very important, because salamander

densities are based on the volume of wood actually searched.

12
Table 4-A hypothetical example of log selectlon In surveys of CWD 

Number encountered 

in decay category 

Log number 

Decay 

category A B C Sample log?= 

1 

2 

3 

4 

5 

6 

7 

8 

9 

10 

11 

12 

13 

14 

15 

16 

17 

18 

19 

20 

and so forthC 

Ab 

A 

C 

B 

B 

A 

A 

: 

A 

C 

A 

:: 

B 

iii 

C 

A 

B 

1 

2 

1 

2 

3 

4 

3 

8 

8 

No 

Yes 

Yes 

No 

No 

No 

No 

Yes 

No 

Yes 

No 

No 

No 

No 

No 

Yes 

No 

Yes 

Yes 

No 

’ Assume that 1 out of every 3 logs is to be sampled, and the following sampling scheme is to be followed: 

category A, log number 2 of 3, category B, log number 3 of 3, and category C, log number 1 of 3. 

b Decay categories: A = decay classes 1 and 2; B = decay class 3; and C = decay classes 4 and 5. 

’ Continue selecting logs until 10 logs in each decay category have been sampled. 

Collecting tips-We have several pointers for more effective collecting for both TCS 

and surveys of CWD. Tools needed for both techniques include potato rakes and 

crowbars. It is necessary to purchase high-quality potato rakes; the less expensive 

ones cannot withstand extensive use. Crowbars are handy for peeling bark and 

breaking up the less-decayed logs. (See appendix 3 for a complete list of materials 

and tools needed to take samples.) 

Large logs and bark piles adjacent to these or large, well-decayed snags are the 

most productive sites for TCS. Follow the instructions above for sampling logs. Cther 

habitats should not be ignored during TCS, however. Moderate-sized debris (10 cm 

or more in diameter) on the forest floor should be turned over; often two people are 

needed to roll logs. In general, avoid raking through leaf litter or turning very small 

objects, but search piles of bark, slash, or mounds, because these often house amphibians. Rocks or boulders, if present, should be turned. Exercise caution when turning rocks on steep slopes. Be alert; searches often occur on rainy days when visibility is poor, especially under closed canopies. Salamanders can flee rapidly down 

13 
a crevice, so grab them by cupping your hand on top of them. Frogs are elusive, and

to catch them you may need the cooperation of two or three people to surround the

quarry. Collectors should scrutinize the area under turned objects. Salamanders often

freeze and most are cryptically colored.

Some species have special traits. Ensatinas are commonly found, and they rarely

move once exposed. They are easily captured but must be picked up carefully or

else they will autotomize (spontaneously amputate) their tail. Newts are slow moving

but possess a highly toxic skin poison. This poison typically is released only if the

animal is under attack but may show up during rough handling (for example, if the

newt is hit by a rake tine). All terrestrial salamanders have some toxic secretions, but

they rarely exude these substances when being handled.

The Oregon slender salamander and the Larch Mountain salamander often coil up,

an apparent mimicry of distasteful millipedes that also curt up. Check any coiled

animal closely. Clouded salamanders and western redback salamanders can move

rapidly and need to be grabbed quickly. At least one hand should be bare to capture

animals; gloves are usually too awkward for collecting agile species.

Snakes might be encountered during searches. Rattlesnakes occur at low elevations

in Oregon and California, especially around rock outcrops. We recommend no collecting of rattlesnakes. Other snakes or lizards can be grabbed or, if fleeing, stepped on

gently. Reptiles should be sluggish in cool, wet weather.

Habitat destruction can be minimized by returning cover items to their original positions. Roll small logs and rocks back and replace large pieces of bark slabs. Rake

decayed logs back together and replace as much bark as possible. Some habitat destruction is unavoidable, but the organic material remains, and at least a portion of

the log-soil interface can be restored by careful replacement of disturbed objects.

Installation of pitfalls-Place pitfall arrays and grids in spots representative of the

study area. If single arrays are to be placed around the study area, the locations

should be preselected from maps or aerial photos. The array or grid location should

be at least 75 m from any forest edges (the farther, the better). For arrays, establish

the center point of the first array at random. If a pair of arrays is used, measure 25 m

from this point in a random direction for the center of the second array. For a grid,

select one corner at random for the location of the grid. The grid is then laid out by

using handheld compasses and 15-m tapes or measured ropes (necessary in dense

brush). Installation of grids is generally fast with a six-person crew; four people lay

out the grid, and two people begin installing traps. Two-person teams are best for

grid layout. One person pulls the tape or rope until stopped by the second person,

who remains at the previous station. Flag the new point and continue.

14
Pitfalls are constructed by fastening the open ends of two number 10 tin cans together with duct tape and then cutting the bottom out of one end (fig. 4). Traps are installed flush with the ground and have a plastic collar inserted at the top. This collar

functions to keep animals from crawling out of the trap and is constructed by cutting

the bottom out of a l-lb plastic margarine tub. When not being used for trapping, the

traps should be closed; use the plastic lids from the margarine tubs. In grids, place

the trap within 2 m of the station flag. If possible, place the trap next to a cover object, such as a rock or downed log. Traps next to logs should be placed on the downhill side of the log. The hole for the trap is dug most easily with a posthole digger,

which creates a hole with the correct diameter. A tile spade can also be used. Traps

have an additional optional wood cover. When the trap is open, the cover is suspended above the opening. This functions in part as a rain cover and partly to attract

animals.

If an array design is being used, drift fences are constructed from 50-cm-tall aluminum valley roofing metal. This comes in 15.2-m rolls, which should be cut into 5-m

sections before it is taken to the study area. We placed fences pointing away from

the center of the array at equal (120o) intervals. The interior end of each fence began

3 m from the center of the array (fig. 3). There are many other possible arrangements

for placing pitfall arrays; see figures in Campbell and Christman (1982), Jones (1981,

1986), and Vogt and Hine (1982).

Figure 4-Construction and placement of a pitfall trap.
Use a mattock or hoe to dig a trench 20 cm deep and 5 m long, stand the fence into

the trench, and back fill with soil. Occasionally an axe is needed to cut large roots.

Tamp down the loose dirt so that the fence is self-supporting (stakes are not necessary for these relatively short fences), and smooth the dirt alongside the fence to

create a runway. Move small obstacles (twigs, rocks) away from the fence. Traps are

placed at the ends of the fence so that no gaps occur between the fence and the rim

of the trap.

There are two important safety rules to follow when installing arrays. First, always

wear gloves to handle the aluminum. The sharp edges can inflict serious cuts on unprotected hands. Second, exercise extreme caution in wet weather. The tools quickly

become coated with slick mud, and a mattock or axe flying out of someone’s hands

is a lethal weapon.

Funnel traps will need to be constructed if reptiles are a target of the study (see

Jones 1986, Vogt and Hine 1982). Funnel traps are constructed from window screen,

which comes in rolls 76 cm wide. Cut a piece 90 cm long, and staple the ends together along the cut edge. Fold back the stapled edge so that you have a tube 25 cm in

diameter by 76 cm long. Construct funnels by rolling square pieces of screen into a

cone and stapling. Fold back the edge and attach to the tube. One end is fastened

permanently with staples, and paper clips are used at the other end so that animals

may be easily removed. Funnel traps are placed midway on both sides of each drift

fence. Shape the trap and fill in with dirt so that no gap occurs between the fence

and the trap. Shade the trap by placing loose bark or litter over the trap.

Pitfall operation-operating pitfall traps is a simple task. Techniques do not differ

between arrays and grids. The primary decision is how frequently the traps should be

checked. Check the traps every other day, if possible, but if there are a large number

of study areas, then traps may have to be checked lessfrequently. Intervals of more

than 5 days between checks should be avoided. Checking traps more frequently produces better specimens, particularly among the mammals that will be caught. If the

number of study sites is such that all traps cannot be opened on the same day, care

must be taken that all traps are closed in the same order they were opened in. This

ensures the same trapping effort for each area.

Each time a trap is checked, remove debris that has fallen into the trap, and bail out

excess water. A small amount of water should be placed in traps when they are

opened, but in wet weather, most traps will accumulate more water than is desired. It

has previously been recommended that water be placed in pitfall traps (Raphael and

Barrett 1981, Williams and Braun 1983), and this is probably the quickest, most humane way to kill small mammals. Current guidelines for using pitfall traps to kill trap

small mammals (American Society of Mammalogists 1987) specify drowning as the

only acceptable method. But drowning is a slow and inhumane way to kill amphibians,

and it has been prohibited in the current guidelines for field methods for herps (ASIH

and others 1987). A generally acceptable compromise between these apparently incompatible recommendations is to keep a small amount of water (2 to 5 cm) in traps

and check them frequently. Small mammals, particularly shrews, will become hypothermic and drown in this amount of water, but most amphibians should be able to

survive.

16
Identification

Disposition of ’

Specimens

All animals trapped in pitfalls are to be returned to the laboratory for processing. Separate mammals, live herps, and dead herps, but otherwise place all animals from the

same trap in one plastic bag. Carry a field notebook with waterproof paper to record

the number of individuals, species, and trap number of all animals caught. This record

is important and should become a permanent part of the data set. It provides critical

information during the initial processing of specimens and is a valuable reference for

the questions that inevitably arise even after the data have been processed. Record

the study area, date, and trap number in pencil on a small piece of waterproof paper

and place in each bag of specimens. Bag all the specimens from a single study area

together in a large plastic bag. Keep the specimens in a cooler with reusable ice containers while in the field. On returning to the lab, place dead specimens in a freezer

and live herps in a cool space or refrigerator.

Accurate identification of specimens in the field is critical for TCS and surveys of

CWD. Field identification is less important for pitfalls, because all specimens are

examined later in the laboratory. The field notes listing the specimens caught in each

trap are more valuable, however, if they are accurate. To increase accuracy, it is

helpful for team members to examine series of specimens at a museum before field

work begins. An additional field practice session is recommended to catch animals

alive and to practice field identification. Most forms have distinct shapes or colors,

but some species present problems. Most people have difficulty with woodland

salamanders (Plethodon spp.), ranid frogs (Rana spp.), and juvenile salamanders.

References for identification of northwestern herps are Nussbaum and others (1983)

and Stebbins (1985). Other useful regional references are by Green and Campbell

(1984) and Gregory and Campbell (1984).

All animals captured in pitfall traps are routinely euthanized and preserved (special

consideration will need to be given to species with special status, such as those listed by the Federal or State governments as threatened or endangered). Specimens

from TCS or CWD surveys may be treated in the same manner, or they may be

released after the surveys near points where they were captured. If specimens are

released, then positive identification is absolutely necessary (see above). Also, if animals are released, a representative series of voucher specimens should be retained

from each study area and preserved. Capturing animals and retaining specimens

requires valid scientific collecting permits from the appropriate State wildlife agency,

and arrangements should be made before the study begins to deposit the specimens

in an appropriate museum.

Process all specimens from a given survey, or all specimens collected from a pitfall

site on a given day, together. This will provide for the most accurate recordkeeping,

and it helps in solving the mystery of the occasional unlabeled specimen. Thaw any

frozen specimens, and kill the live ones. Be sure to keep the label identifying the

specimen closely associated with each specimen. Kill by relaxing amphibians in a

dilute solution of Chloretone and by injecting reptiles with aqueous sodium pentobarbital. Chloretone is a saturated solution of hydrous chlorobutanol in 95 percent

17
ethanol. An effective dilution is 2 ml per 570 ml of water. Sodium pentobarbital

(Nembuto12 is one trade name) is a restricted drug and may be difficult to obtain.

Reptiles may also be killed by injecting 95 percent ethanol into the heart region.

After the animal is dead, weigh and measure it (see appendix 2), tie a numbered

tag to the right hind leg, and preserve in formalin. Create a lo-percent solution of

buffered formalin by diluting commercial formalin to 10 percent and adding 4 g of

baking soda or sodium carbonate per 400 ml of solution. Amphibians that appeared

dead may begin to move when placed in the formalin. These should immediately be

rinsed in water and returned to the Chloretone until dead. Amphibians and lizards

should be laid out ventral side down in a shallow pan with a tight-fitting lid; for

example, a plastic freezer container. Line the bottom of the pan with commercial

paper towels (household towels have “dimples” that become imprinted on the skins

of the animals), and pour a small amount of formalin into the pan. Snakes should be

folded into an oblong coil with the head on the inside. The coil should be short

enough to fit in the storage jars. Reptiles also must have formalin injected into the

body cavity, limbs, and tail. Do not inject so much that a balloonlike specimen is

created. If injection is not possible, then the body cavity, limbs, and tail must be slit

to allow the formalin to enter the body. Body cavities of large Pacific giant salamanders should also be slit for thorough preservation. Pisani (1973) provides a thorough

discussion of preservation techniques. Let the specimens fix in the formalin for at

least 24 hours, then store in 50 percent isopropyl alcohol.

If specimens are released, then reasonably accurate measures of snout-vent and

total lengths can still be made. Place the animal in a plastic bag and restrain it

against the bottom of the bag. When the animal is quiet and relatively straight,

measure to the nearest millimeter with a ruler. Mass can also be measured in the

field with spring scales available in forestry supply catalogs.

The investigator should be aware that in northwestern forests, twice as many small

mammals as herps generally are captured in pitfall traps. If a study is planned that

uses pitfall traps, provision should be made for preserving the mammals. Neglecting

this would be a criminal waste of valuable data.

Data Analysis Numerous analyses can be done on the types of data collected from surveys of

amphibian occurrence and abundance (see papers in Szaro and others [1988] and

Ruggiero and others [in press] for examples). We will give a couple examples of the

types of analyses that can be done, and we will discuss any special analyses that

need to be performed.

All the techniques are excellent at providing data on presence or absence of species,

and two or more techniques can be combined to provide a complete assessment of

all the species potentially present. One example is provided by considering amphibians and reptiles detected by pitfall trapping with arrays and TCS at 18 study areas

2

 The use of trade, firm, or corporation names in this publication is for the information and convenience of the reader.

Such use does not constitute an official endorsement or approval by the U.S. Department of Agriculture of any product

or service to the exclusion of others that may be suitable.

18
in the Oregon Cascade Range in 1983 (table 5). Presence-absence data can be analyzed by calculating measures of similarity and then using a clustering procedure to

look for patterns among groups of study areas (Pielou 1984). From the data matrix in

table 5, similarities were calculated for every pair of stands by using Jaccard’s index

(Pielou 1984), which is the percentage of species both areas have in common compared to the total number of species present at either area. Clustering was accomplished by using the nearest-neighbor technique. One group of five old-growth and

mature stands cluster together above the 60-percent level of similarity, but in general,

there are few recognizable patterns related to habitat type (fig. 5). Pielou (1984) and

Gauch (1982) are valuable sources of techniques for analyzing the structure and

organization of communities.

Table 5-List of species of amphibians and reptiles present (P) at 18 study areas in and near

the H.J. Andrews Experimental Forest, Lane County, Oregon, 1983

Stand number

Old-growth stands Mature stands Young stands Clearcut stands

Species 2 3 15 17 24 25 29 33 11 35 39 42 47 40 75 55 92 93

Northwestern salamander

Pacific giant salamander

Clouded salamander

Oregon slender salamander

Ensatina

Dunn’s salamander

Rough-skinned newt

Tailed frog

Pacific trecfrog

Red-legged frog

Reptiles:

Western skink

Northern alligator lizard

Western fence lizard

Rubber boa

Northwestern garter snake

Common garter snake

P P

P P

P P P P P P

P P P P P

P P P P P P P

P P P

P P P P P P

P P P P P

P P

P

P

P P P

P

P P P

P P P

,;lj c’

‘.. I.

P P

P P

P P P

P

P P

P

P

P P

P P P

P P P P

P P P P

P P P P P

P P P P P P

P P P P P

P P P

P P P

-P

P

P

P

P

P

P P

P P

P P

P

P P

P P

P

P P

P

.-

P

P _. P

P

P

P

Number of species 8 5 6 7 6 7 11 3 5 6 4 6 5 8 9 9 13 6

19
slMllARlTv 

100 80 50 40 20 0 

I I I I I I I I I I STANDTYPE # I 

OLDQROWlli 24 

c4DQrn 3 

MAlulE 11 

uDQRowM 17 

OIDQ~ 2 

ct.- 55 

MATURE 35 

YWNQ 75 

OLDQROWTH 29 

92 

Ot.DQm 15 

YOUNQ 40 

OLDQf3OWlH 33 

Figure 5-Nearest-neighbor clustering of herpetofauna at 18 study areas in the Oregon Cascade 

Range in 1983. Similarities were calculated by using presence-absence combined data from 

pitfall trapping and TCS. 

.ei?) 

Surveys of CWD can provide initial estimates of population density. The density in 

downed wood of each species of salamander (number per cubic meter) is calculated 

as the number caught in each log, divided by the volume of wood sampled in each 

log. Mean densities in downed wood in each stand were calculated for each of the 

three decay categories (decay classes 1 and 2, class 3, and classes 4 and 5). Use a 

nested analysis of variance (stands within forest age classes) to test whether density 

(log transformed) in downed wood of any species varies among decay categories or 

age class (old growth, mature, and young growth). 

We calculated predicted densities of plethodontid salamanders in 45 forest stands 

from the following formula: 

3 

D = C (di*Vi) , 

i=l 

where D = number of salamanders per ha, di = density in downed wood in decay 

category i, and Vi = m3 of downed wood per ha in category i. See Spies and others 

(1988) for techniques to determine the amount of downed wood present in a stand. 

Where d varied among age classes, D was calculated by using the mean density in 

downed wood for each age class. 

20 
-

l

; . STAND AGE (YEARS)

Figure B-Estimated densities of salamanders relative to forest age

in the Oregon Coast Ranges in 1985. Data are based on surveys

of CWD at 15 study areas. Density was estimated for an additional

30 stands, for which data existed on the volume of downed wood

present in the study area.

Estimated density of plethodontid salamanders was related to stand age for 45 study

areas in the Oregon Coast Ranges in 1985 (fig. 6). There were 15 study areas with

surveys of CWD. Densities in the remaining 36 areas were estimated by using the

average values of d for each habitat type and the measured value of V for each area.

Conclusions There is a vast literature on techniques for sampling and analyzing vertebrate populations, but it was not our intention to provide a complete overview. Rather, we have

described the specialized methods for sampling herpetofauna that we have used and

refined in 3 years of field work in the forests of Oregon and Washington. Comprehensive references on sampling techniques include Cooperrider and others (1986) and

Schemnitz (1980).

Acknowledgments

The methods we have described are most appropriate for surveys of forest-dwelling

amphibians. Because these species use several habitats for breeding, feeding, and

cover and differ widely in vagility, no single method is adequate to sample the entire

community. Pitfall trapping needs to be combined with either time-constrained collecting or surveys of coarse woody debris in any comprehensive survey. _.

We thank Michael Bogan, Andrew Carey, Lawrence Jones, and Leonard Ruggiero for

reading and commenting on this paper. Development of these techniques was aided

by Lawrence Jones and Martin Raphael and a dedicated corps of field biologists. This

is contribution number 68 of the Wildlife Habitat Relationships in Western Oregon

and Washington Project.

21
When you know: Multiply by: To find:

Centimeters (cm)

Meters (m)

Square meters (m2)

Hectares (ha)

Cubic meters (m3)

Grams (g)

Milliliters (ml)

0.394 Inches

3.281 Feet

10.764 Square feet

2.471 Acres

35.315 Cubic feet

0.035 Ounces

0.035 Fluid ounces

American Society of Ichthyologists and Herpetologists; The Herpetologists’

League; Society for the Study of Amphibians and Reptiles. 1987. Guidelines

for the use of live amphibians and reptiles in field research. [Location of publishers

unknown]: 14 p.

American Society of Mammaloglsts. 1987. Acceptible field methods in mammalogy: preliminary guidelines approved by the American Society of Mammalogists.

Journal of Mammalogy. 68 (suppl.): l-18.

Aubry, Keith B.; Jones, Lawrence L.C.; Hall, Patricia A. 1988. Use of woody

debris by plethodontid salamanders in Douglas-fir in Washington. In: Szaro,

Robert C.; Severson, Keith E.; Patton, David R., tech. coords. Management of

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Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain

Forest and Range Experiment Station: 32-37.

Banks, Richard C.; McDiarmid, Roy W.; Gardner, Alfred L. 1987. Checklist of

vertebrates of the United States, the U. S. Territories, and Canada. Resour. Publ.

166. Washington, DC: U.S. Department of the Interior, Fish and Wildlife Service.

79 p.

Burton, Thomas M.; Likens, Gene E. 1975. Salamander populations and biomass

in the Hubbard Brook Experimental Forest, New Hampshire. Copeia. 1975:

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Bury, R. Bruce. 1983. Differences in amphibian populations in logged and old growth

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Bury, R. Bruce. 1988. Habitat relationships and ecological importance of amphibians

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University of Washington, Seattle. Contribution 59. Seattle: Institute of Forest

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22
Bury, R. Bruce; Corn, Paul Stephen. 1988. Douglas-fir forests in the Oregon and

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and monitoring of wildlife habitat. Denver, CO: U.S. Department of the Interior,

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in the Oregon Coast Ranges. In: Ruggiero, Leonard F.; Aubry, Keith B.; Carey,

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Pacific Northwest Research Station. ,

,.

Franklin, J.F.; Cromack, K., Jr.; Denison, W. [and others]. 1981. Ecological

characteristics of old-growth Douglas-fir forests. Gen. Tech. Rep. PNW-118. ”

Portland, OR: U.S. Department of Agriculture, Forest Service, Pacific Northwest

Forest and Range Experiment Station. 48 p.

Friend, Gordon R. 1984. Relative efficiency of two pitfall-drift fence systems for

sampling small vertebrates. Australian Zoologist. 21: 423-433.

23
Gauch, Hugh G., Jr. 1982. Multivariate analysis in community ecology. New York:

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.:rj:;

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24
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25
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26
Appendix 1 Table 6-Scientific and common names of amphibians and reptiles found in the

Pacific Northwest west of the Cascade Range from northern California to

British Columbia

Scientific namea

4-letter

code Common namea

Amphibia, order Urodela (salamanders):

Family AmbystomatidaeAmbystoma gracile

A. macrodactylum

Family DicamptodontidaeDicamptodon copei

D. ensatus

D. tenebrosus

Rhyacotriton olmpicus

Family PlethodontidaeAneides ferreus

A. flavipunctatus

A. lugubris

Batrachoseps attenuatus

B. wrighti

Plethodon dunni

P. elongatus

AMGR

AMMA

DICO

DIEN

DITE

RHOL

ANFE

ANFL

ANLU

BAAT

BAWR

PLDU

PLEL

P, larselli PLLA

P. vandykei PLVA

P. vehiculum PLVE

Family SalamandridaeTaricha granulosa

T. rivularis

T. torosa

Amphibia, order Anura (frogs and toads):

Family Leiopelmatidae, Ascaphus truei

Family Bufonidae, Bufo boreas

Family Hylidae, Hyla regilla

Family RanidaeRana aurora

R. boylii

R. cascadae

R. catesbeiana

R. clamitans

R. pretiosa

Reptilia, order Chelonia (turtles):

Family EmydidaeChrysemys picta

Clemmys marmorata

Reptilia, order Squamata (lizards and snakes):

Family AnguidaeGerrhonotus coeruleus

G. multicarinatus

Family IguanidaePhrynosoma douglassii

Sceloporus graciosus

S. occidentalis

Family Scincidae, Eumeces skiltonianus

Family Boidae, Charina bottae

TAGR

TARI

TATO

ASTR

BUBO

HYRE

RAAU

RABO

RACA

RACT

RACL

RAPR

CHPI

CLMA

GECO

GEMU

PHDO

SCGR

scoc

EUSK

CHBO

Northwestern salamander

Long-toed salamander

Cope’s giant salamander

California giant salamander

Pacific giant salamander

Olympic salamander

Clouded salamander

Black salamander

Arboreal salamander

California slender salamander

Oregon slender salamander

Dunn’s salamander

Del Norte salamander

(includes P, stormi)

Larch Mountain salamander

Van Dyke’s Salamander

Western red-backed

salamander

Rough-skinned newt

Red-bellied newt

California newt

Tailed frog

Western toad

Pacific treefrog

Red-legged frog

Foothill yellow-legged frog

Cascades frog

Bullfrog (introduced)

Green frog (introduced)

Spotted frog

Painted turtle

Western pond turtle

-.

Northern alligator lizard

Southern alligator lizard

Short-horned lizard

Sagebrush lizard

Western fence lizard

Western skink

Rubber boa

27
Appendix 2

Data Sheets

28

Table 6-continued

4-letter

Scientific nameacode

Family ColubridaeColuber constrictor coca

Contia tenuis COTE

Diadophis punctatus DIPU

Lampropeltis getulus LAGE

L. zonata LAZO

Masticophis taeniatus MATE

Pituophis melanoleucus PIME

Thamnophis couchi THCO

T. elegans THEL

T. ordinoides THOR

T. sirtalis THSI

Family Crotalidae, Crotalus viridis CRVI

a

 Scientific and common names follow Banks and others (1987).

Sources: Nussbaum and others (1983) and Stebbins (1985).

Common name’

Racer

Sharptail snake

Ringneck snake

Common king snake

California mountain kingsnake

Striped whipsnake

Gopher snake

Sierra garter snake

Western terrestrial garter snake

Northwestern garter snake

Common garter snake

Western rattlesnake

Data sheet for TCS-This data sheet (fig. 7) needs to be on waterproof paper. The

number of data sheets needed will depend on the number of animals captured. Note

that each area search (1 staff hour) is listed separately and there is room for five

animals per search. If more than five animals are captured in one area, then continue

the data in the space for the next area, but if fewer than five animals are captured,

then skip to the space for the next area before recording data from the new area.

Data categories are explained below:

1. Standard header. This will differ by study. We illustrate the information we

recorded in the old-growth study.

2. Weather (WR). Use the codes listed at the bottom of the data sheet.

3. Temperature (

oC).

4. Start time, end time. Use 24-hour notation.

5. Crew. List the initials of the other crew members. The recorder should be the

same person for each stand. Note whether or not the recorder participated in

the collecting.

6. Catalog initials. Initials of the collector in whose catalog the specimens will be

recorded.

7. Area. Each-l -staff hour search should be numbered sequentially.

8. Aspect (degrees). Record for each area searched.

9. Slope (percent). Record for each area searched.

10. Specimen number. Each herptile encountered is given a unique number, either

sequentially for the entire stand (1, 2, 3,...n), or sequentially for each area

searched (l-l, 1-2, l-3,...n; 2-1, 2-2, and so forth). Whichever method is used,

the data collected in the laboratory (see below) must be matched to the data

collected in the field.

11. Catalog number. This is the number given to preserved specimens. We use

small, rectangular tags, preprinted with the catalog number.

12. Species. This is the four-digit code for each species (see appendix 1).
TIME-CONSTRAtNED COLLECTING DATA 

snout-vent total W&A rree 

;Tpec# II 

d-v 

catalog number ,, 

cove

SpCieS ..= ..lagth (mm) ..lw@ (mm) __ m= 0 ~0s SpWkS *e= *. leJ%o 

snout-vent mee decay cove

VERTICAL FOSlTlON: WEATHER 

l-onsulface 7 - Under Bark on Log 

2 - Under Rock (not in talus) 

3-UnderRockinTalus 

8-UnderBarkonGround~ ImfromSnag 

4-UnderlBg 

9-UnderBarkonGround < I mfromLog 

10 - Under Bark on Ground 

5-InlDg 

6-UnderE~arkonSnag ’ 

(>lmfromlogorsnag) 

11 -other 

l-clear 

2-Paltlycloudy 

3-overcast 

4-Fogonly 

5-Dride 

6 - S

7-Ra

8 - S

Figure 7-Data sheet for recording information from time-constrained surveys. This sheet should be on 

waterproof paper. 
The following data items (13-17) are recorded most accurately from anesthetized

animals in the laboratory. Animals should be placed individually in plastic bags with

the specimen number (item 9) so that the data can be properly recorded. If animals

are released after collecting, these data can still be recorded; they will have slightly

lower accuracy.

13. Sex. M = male; F = female; if unknown, leave blank.

14. Age. A = adult, J = juvenile.

15. Snout-vent length. Record to the nearest 0.1 millimeter.

16. Total length. Record to the nearest 0.1 millimeter. If the tail has been broken or

is otherwise incomplete, leave this blank.

17. Mass. Record to the nearest 0.1 gram.

Items 18-21 are recorded in the field.

18. Vertical position. Use the codes at the bottom of the data sheet.

19. Tree species. The four-digit code for the species (if known) of the snag or log.

20. Decay class. Use the code for either snags or logs, as appropriate.

21. Cover-object dimensions. Record length and width to the nearest centimeter.

Data sheet for surveys of CWD-This data sheet (fig. 8) also needs to be on waterproof paper. Data for the specimens collected at each log are recorded directly below

the data for the log. At least 15 data sheets will be needed per study area. As with

the data for TCS, there is room for five animals per log. If the number of animals captured exceeds the space available, then follow the same procedures as for TCS.

Data items are explained below:

1.

2.

3.

4.

5.

6.

7.

8.

9.

Standard header. The first two lines at the top of the page are the same as for

TCS. The following items (2-13) are data collected on each log before it is

searched for animals.

Log number. Number logs sequentially from the start of each survey.

Time. Record the number of minutes required to search the log (20 staff minutes,

maximum).

Decay class. Use the five-class scale. Other decay categories can be assigned

during data analysis.

Tree species.

Aspect.

Slope. Record the percent slope over a 10-m run, with the log at the midpoint.

Total log: length. Record to the nearest meter.

Total log: maximum diameter (cm).

10. Total log: minimum diameter (cm).

11. Portion sampled: length (m). Record the amount of the log that was actually

searched.

12. Portion sampled: maximum diameter (cm).

13. Portion sampled: minimum diameter (cm).

30
LOG SURVEY DATA 

technique province 

JI-llday-l--J&t 

habitat stand # month year temp start time end time crew 

II-II 

uamn-n~ffimmommi3mnmm

d-y 

total log: portio

1% # , 

time class ,, , tree species aspect % slope length (m) ,max diam (cm) , min diam (cm) length (m) length (m) ,max diam (cm) , min diam (cm) length (m) max 

mmnmnmmim3iin3m2~ 

so&menpecimen posi- odeathp snout-vent total 

d-y 

log # , time ,, , class aspect 

total log: portion 

% slope length (m) max diam (cm) min diam (cm) length (m) max d

posi- depth snout-vent 

catalog number 

W 

2 POSITION: 1 - Under Bark (on log); 2 - Under Bark (on ground); 3 - Under Log; 4 - In Log; 5 - Other 

Figure 8-Data sheet for recording information from surveys of coarse woody debris. This sheet should be on waterproof paper, 
The following items are collected for each animal encountered. Most are the same as

for TCS and may be recorded in the field or in the lab, if all the animals are retained.

Data unique to log surveys that are recorded in the field are:

14. Position (POS). Use the codes at the bottom of the data sheet.

15. Depth in log (cm). Record the distance to the exterior surface of the log.

Pitfall trapping data sheet-These data (fig. 9) are recorded in the lab when the

animals are processed. A waterproof sheet is not necessary. Use a new data sheet

for each time the traps are checked. Most of the data are the same as those

collected for TCS and surveys of CWD. Unique elements are:

1. Trap night. Record the number of nights since the traps were opened; for

example, if the traps were opened October 1, and these are data for animals

picked up when the traps were checked on October 18, then trap nights are 17.

2. Trap number. Record the trap position (column and row) for each animal.

32
PITFALL DATA 

mP snout-vent 

catalog number ,, species . . length (mm) 

TRAP NIGHT: Number of nigh& since the traps were opened. 

SEX: Male, &male (if unknown, leave blank) 

AGE: &iult, Subadult, Juvenile 

Figure g--Data sheet for recording information on animals collected in pitfall traps. This sheet does not need to be on waterproof paper 

33 
Appendix 3 Item

Materials Needed for

TCS or Surveys of CWD

Materials Needed for

Pitfall Installation

and Operation

Materials Needed

in the Lab

Number

Potato rakes (a backup rake is not a bad idea)

Crowbar

Stopwatch

Clipboard

Thermometer

Plastic bags

Cloth bags or pillowcases

Pencils

Compass

Clinometer

Short (15 cm) plastic ruler

Long (30 cm) plastic ruler

10-m measuring tape

Item Number

2

1

1

1

1

several

1 or2

2+

1

1

1

1

1

Installation

Posthole digger (l/person)

15-m tape or measured nylon rope

Plastic flagging (1 roll/pair of people)

Waterproof ink marker (l/person)

Number 10 tin cans

l-lb margarine tubs

Wood covers

Operation

Waterproof notebook and paper (l/person)

6- by 10-inch plastic bags

12- by 16-inch plastic bags

Plastic cup or long handled spoon (l/person)

Small cooler with reusable refrigerant

Calipers

Plastic ruler (30 cm)

Spring scales (10 g, 50 g, 100 g)

Scissors

Forceps

Tags with preprinted catalog numbers

Paper towels (industrial type)

Plastic trays with lids

Cloretone

Nembutol

Formalin (40 percent formaldehyde solution)

95 percent ethanol

Isopropyl alcohol (dilute to 50 percent)

Jars for specimen storage

I+

2+

l +

l +

72/grid, or

24/array

36/grid, of

12/array

36/grid, or

12/array

l +

Many

Many

 l+

l +

34