Vancouver Island AmphibiansT. M. Davis and A. Dehalt

Amphibians are ectothermic vertebrates with glandular skin lacking any epidermal coverings such as fur, scales or feathers. Like mammals, they have two occipital condyles on the skull (unlike like reptiles and birds that have only one). Their anamniotic eggs lack shells and extra-embryonic membranes, so they must be laid in water or moist environments. Class Amphibia contains over 6,500 extant species in three orders: order Anura (frogs and toads), order Caudata (salamanders and newts) and order Gymnophiona (caecilians - legless amphibians).

Toads are frogs with stubby bodies, short hind legs (for walking instead of hopping), warts and dry skin, and have parotoid (poison) glands behind the eyes. Newts are aquatic or semi-aquatic salamanders in Family Salamandridae and lack costal grooves .

Amphibians of Vancouver Island

Class AmphibiaOrder Caudata (salamanders)

Family Ambystomatidae (mole salamanders)Long-toed Salamander (Ambystoma macrodactylum) - AMMANorthwestern Salamander (Ambystoma gracile) - AMGR

Family Salamandridae (newts)Rough-skinned Newt (Taricha granulosa) - TAGR

Family Plethodontidae (lungless salamanders)Ensatina (Ensatina eschscholtzii) - ENESWandering Salamander (Aneides vagrans) - ANVAWestern Red-backed Salamander (Plethodon vehiculum) - PLVE

Order Anura (frogs and toads)Family Bufonidae (true toads)

Western Toad (Anaxyrus boreas) - ANBO

Family Hylidae (treefrogs)Northern Pacific Tree (aka “Chorus”) Frog (Pseudacris regilla) – PSRE

Family Ranidae (true frogs)

Red-legged Frog (Rana aurora) - RAAUGreen Frog (Lithobates clamitans)* – LICLAmerican Bullfrog (Lithobates catesbeiana)* – LICA

*introduced species

VANCOUVER ISLAND AMPHIBIANS

Key to the Terrestrial Salamanders of Vancouver Island(modified from Matsuda et al. 2006)

1a. Skin rough and tuberculate; belly bright orange; no obvious costal groves ... ………………………………………………Rough-skinned Newt (Taricha granulosa)

1b. Skin smooth; costal grooves evident 2

2a. Naso-labial groove running down from each nostril; parasphenoid teeth present.... 3

2a. No naso-labial grooves or parasphenoid teeth present 6

3a. Smooth-edged yellow, orange or red dorsal stripe continuing to tip of tail ... …………………………….Western Red-backed Salamander (Plethodon vehiculum)

3b. No dorsal stripe 4

4a. Tail constricted at base; 12 costal grooves on each side of body .... ……………………………………………………… Ensatina (Ensatina eschscholtzii)

4a. Tail not constricted at base; 15 costal grooves on each side of body 5

5a. Square-tipped toes; mottled grey color, but sub-adults often black and juveniles with copper triangle on snout Wandering Salamander (Aneides vagrans)

5b. Toe tips rounded; uniform grey to black ... ……...melanistic morph of Western Red-backed Salamander (Plethodon vehiculum)

6a. Irregular green or yellow dorsal strip breaking up on the head and tail; white speckleson sides and belly; second toe from outside on each hind foot longer than other toes ...……………………………………Long-toed Salamander (Ambystoma macrodactylum)

6b. No dorsal stripe; dark brown to black or grey; length more than 120 mm; distinct paratoid gland present ...........................Northwestern Salamander (Ambystoma gracile)

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Key to the Adult Frogs and Toads of Vancouver Island (modified from Matsuda et al. 2006)

1a. Parotoid glands and rough warts on skin; body squat ...………………………………………………………...Western Toad (Anaxyrus boreas)

1b. No parotoid glands or warts; skin smooth 2

2a. Adhesive toe pads conspicuous; dark eye stripe ending at shoulder ...………………………………………..Northern Pacific Tree Frog (Pseudacris regilla)

2b. Toe pads absent or inconspicuous 3

3a. Brown or tan; obvious dark facial mask ..................Red-legged Frog (Rana aurora)

3b. Green or olive-green; no dark facial mask 4

4a. Dorso-lateral folds present ................................... Green Frog (Lithobates clamitans)

4b. Dorso-lateral folds absent; skin fold around ear membrane present ... ……………………………………….……American Bullfrog (Lithobates catesbeiana)

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Sampling and Monitoring AmphibiansHerpetology is the study of amphibians and (non-avian) reptiles. Because most species in these two classes are typically small, secretive and inconspicuous, they are often thought to be relatively unimportant ecologically. However, they are good bio-indicators of the functional status of an ecosystem, and their biomass can exceed that of other vertebrates in the same area. Unfortunately, there have been numerous declines and extinctions of amphibian and reptile populations recently in many parts of the world, primarily as the result human activities that fragment, pollute, or destroy critical habitats.

Determining the abundance of amphibians and reptiles is important with respect to understanding their ecology and is critical for conservation efforts. However, naive application of sampling methods without knowledge of the organism’s natural history will likely produce spurious results. Also, management decisions should not be based on simple or superficial measures of biodiversity, but instead on detailed knowledge of the ecology and behaviour of individual species, and this can only be obtained by intensive in-depth studies of local populations.

In B.C. there are relatively few species of reptiles and amphibians, but they are tremendously diverse in terms of morphology, life history, ecology and behaviour. Because of this diversity, sampling different species within an area often requires a variety of sampling methods. The number of individuals and the particular species one finds on random searches depends on a variety of physical, temporal, biological, and other factors. Sampling methods include call indices (frogs), pitfall and funnel traps, artificial cover objects (ACO), transect searches of natural cover, nocturnal road cruising and other methods (see Heyer et al. 1994 for a review).

Special methods for sampling amphibians in streams and lakes have been developed (Bury and Corn 1991; Olsen et al. 1997). Standard methods for determining terrestrial salamander abundance include mark-and-recapture, time-constrained searches (TCS), area-constrained searches (ACS), surveys of coarse woody debris (CWD), cover boards and pitfall traps. However, serious errors can arise if these methods are used uncritically.

The objectives of most inventories are to either determine 1) presence/not detected (what species occur in a particular area) or 2) relative abundance (a measure of the number of organisms at one location or time relative to the number of organisms at another location or time). Sometimes, researchers are interested in determining absolute abundance (the actual total population size), but this is labour-intensive and methodologically complex.

Although “presence” can be established unambiguously, “absence” is problematic. “Absence” is inferred by repeated searches that fail to find the target species. However, that a species was not found on repeated searches does not guarantee that it is not present. Rare, cryptic and secretive species are often difficult to detect and apparent absence can be the result of temporary, local extinction or a lack of breeding or delayed breeding in a particular year. Most researchers therefore prefer “not detected” to “absence” because of this uncertainty (BC Wildlife Federation 2002). However, if the appropriate microhabitats are searched when weather conditions are favorable and searches are done several times in a season, it becomes increasing unlikely that the species is present as the cumulative number of unproductive searches increases.

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Relative abundance is the most commonly used measure of amphibian abundance. It can be calculated in reference to either area or time/effort (e.g. number of person-hours of searching). We will use an area-constrained search (ACS) of 1-m2 quadrats to search for Plethodontid salamanders and opportunistic time-constrained searches (TCS) to search for both salamanders and anurans (frogs and toads). Time permitting, we may also do some night road cruising.

Area-constrained Search (ACS)We will search 1-m2 quadrats placed randomly along equally spaced transects. The advantage of equally-spaced transects is that they are convenient. Randomness with respect to habitat is achieved by spacing the quadrats randomly along the transects. The group will be divided in to teams of 3 (or 4) members and each team will search 5 quadrats along their own transect. Your instructor will determine the starting position (based on previous sampling) of the first transect and subsequent groups will start their transects 5 m from the adjacent transect. Transects will be parallel to each other (follow an agreed upon compass reading) and perpendicular to the road. To avoid edge effect, start your transect 10 m from the edge of the road.

To determine the position of the five quadrats along your transect, select five numbers between 2 and 10 from a table of random numbers (Tab.1) or by any other unbiased method. This will be the number of meters between quadrats, measured from the far edge of the previous quadrat. If your quadrat falls on a large log or rock that you cannot move, move the quadrat to just the other (far) side of the log or rock. Otherwise, however, do not pick “better” or more convenient spots for your quadrats because this would undermine the whole point of randomization (which is to sample without bias).

Follow the following procedures on your search:

1. At your sample point, set down your quadrat frame. Be sure not to step in the quadrat.

2. Begin by searching the perimeter of the quadrat and work in towards the center (Fig.1). This avoids having salamanders escape from the quadrat before they are discovered.

Figure 1. Searching a 1x1 m quadrat: outside-in.

3. Remove all debris (rocks, pieces of bark or wood, and leaves) from the quadrat.

4. Place any amphibians in its own zip-lock bag with a bit of moss or soil for moisture. Measure and weigh the animals only after the entire quadrat has been searched.

5. Replace all debris once the quadrat search is complete.

6. Release the amphibians back into the quadrat.

7. Estimate the surface density of each species by taking the mean of all quadrats.

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RANDOM NUMBER TABLE

20 17

42 28

23 17

59 66

38 61

02 10

86 10

51 55

92 52

44 25

74 49

04 19

03 04

10 33

53 70

11 54

48 63

94 60

94 49

57 38

94 70

49 31

38 67

23 42

29 65

40 88

78 71

37 18

48 64

06 57

22 15

78 15

69 84

32 52

32 54

15 12

54 02

01 37

38 37

12 93

93 29

12 18

27 30

30 55

91 87

50 57

58 51

49 36

12 53

96 40

45 04

77 97

36 14

99 45

52 95

69 85

03 83

51 87

85 56

22 37

44 91

99 49

89 39

94 60

48 49

06 77

64 72

59 26

08 51

25 57

16 23

91 02

19 96

47 59

89 65

27 84

30 92

63 37

26 24

23 66

04 50

65 04

65 65

82 42

70 51

55 04

61 47

88 83

99 34

82 37

32 70

17 72

03 61

66 26

24 71

22 77

88 33

17 78

08 92

73 49

03 64

59 07

42 95

81 39

06 41

20 81

92 34

51 90

39 08

21 42

62 49

00 90

67 86

93 48

31 83

19 07

67 68

49 03

27 47

52 03

61 00

95 86

98 36

14 03

48 88

51 07

33 40

06 86

33 76

68 57

89 03

90 49

28 74

21 04

09 96

60 45

22 03

52 80

01 79

33 81

01 72

33 85

52 40

60 07

06 71

89 27

14 29

55 24

85 79

31 96

27 56

49 79

34 34

32 22

60 53

91 17

33 26

44 70

93 14

99 70

49 05

74 48

10 55

35 25

24 28

20 22

35 66

66 34

26 35

91 23

49 74

37 25

97 26

33 94

42 23

01 28

59 58

92 69

03 66

73 82

Tab. 1 Source: "Statistics for Biologists by R.C. Campbell, Cambridge University Press, 1974.

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If randomly picking a number between 2 and 10, you need to decide beforehand which digit(s) to use of each 4 or 5-digit number (depending on given table), how to differentiate between 1 and 10, whether to pick each number separately or to proceed up, down, left, right from the first randomly picked number, etc.

Time-constrained search (TCS)

A time-constrained search (TCS) involves searching an area for a specified amount of time and can be used for both presence/not detected sampling and determining relative abundance. This is an equal-effort search as measured by the number of person-hours spent searching (Corn and Bury 1990). A TCS should cover as much habitat as possible, so no more than a few minutes should be spent searching any one object.

Care must be taken not to search where other team members have searched, so an informal systemic approach is appropriate. Search the habitat by turning over cover objects (logs, lark on the ground, rocks) and searching species-specific microhabitats. For example, Aneides vagrans will most likely be found by carefully peeling off the bark on longs in a mid-state stage of decay. The bark should be replaced in the same position as it was found.

The time searching should be kept independently by each searcher and then summed for the whole group. Do not count time measuring or weighing or writing notes; only count time actually searching (which includes time walking from cover object to cover object).

Handling and Measuring Amphibians

You should always be careful not to harm live amphibians while examining them. Frogs and salamanders, especially hatchlings and juveniles, are highly sensitive and delicate animals and should only be handled when necessary. All amphibians are ectotherms and are less active at colder temperatures. Thus, it is easier to take measurements when the weather is cold because they struggle less and are easier to capture. Also, cold hands are better than warm hands when handling salamanders for the same reason.

Because of their non-keratinized skin (no fur, feathers, scales…), amphibians are subject to desiccation, they can very quickly dehydrate when handling them. Therefore, handling should be kept to a minimum and a spray bottle with untreated water should be kept handy.

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Here are some basic rules to follow:

Make sure your hands are wet, clean, and free of insect repellent, suntan lotion, or toxins (or potential disease) from a previously handled amphibian.

If a frog or toad appears ill, handle it with disposable gloves to avoid transfer of diseases to other individuals. Where disease is suspected, handle each individual with a new pair of disposal gloves.

Be careful not to let amphibians get overheated or dried out. Spray with untreated water from the nearest natural (uncontaminated) fresh water source if necessary. Let the animal rest in a container.

Because Plethodontid salamanders are lungless, they soon run out of oxygen when struggling and will become limp. They will, however, revive after a short rest, but you should avoid stressing them to that extent.

To examine hatchlings, larval salamanders and tadpoles, put them in a clear plastic container.

Adult salamanders must be kept cool and if you hand is warm, the salamander might become agitated.

Do not detach individual eggs from egg masses and do not detach egg masses from vegetation.

Do not catch a salamander by its tail. It might break off!

Frogs and toads are easier to hold if you let the animal extend its hind legs, and then you gently hold the legs together while you support the body.

Clean and disinfect all field equipment after each use. To prevent the transfer of diseases, fungi, seeds and other unwanted organisms from one site to another, make sure nets, boots plastic bags and all other equipment is washed and disinfected with alcohol or bleach before moving to a new area (see DAPTF guidelines below).

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The Declining Amphibian Populations Task Force Fieldwork Code of Practice

A code of practice, prepared by the Declining Amphibian Populations Task Force (DAPTF) to provide guidelines for use by anyone conducting field work at amphibian breeding sites or in other aquatic habitats. Observations of diseased and parasite-infected amphibians are now being frequently reported from sites all over the world. This has given rise to concerns that releasing amphibians following a period of captivity, during which time they can pick up unapparent infections of novel disease agents, may cause an increased risk of mortality in wild populations. Amphibian pathogens and parasites can also be carried in a variety of ways between habitats on the hands, footwear, or equipment of fieldworkers, which can spread them to novel localities containing species which have had little or no prior contact with such pathogens or parasites. Such occurrences may be implicated in some instances where amphibian populations have declined. Therefore, it is vitally important for those involved in amphibian research (and other wetland/pond studies including those on fish, invertebrates and plants) to take steps to minimize the spread of disease and parasites between study sites.

1. Remove mud, snails, algae, and other debris from nets, traps, boots, vehicle tires and all other surfaces. Rinse cleaned items with sterilized (e.g. boiled or treated) water before leaving each study site.

2. Boots, nets, traps, etc., should then be scrubbed with 70% ethanol solution (or sodium hypochlorite 3 to 6%) and rinsed clean with sterilized water between study sites. Avoid cleaning equipment in the immediate vicinity of a pond or wetland.

3. In remote locations, clean all equipment as described above upon return to the lab or "base camp". Elsewhere, when washing machine facilities are available, remove nets from poles and wash with bleach on a "delicates" cycle, contained in a protective mesh laundry bag.

4. When working at sites with known or suspected disease problems, or when sampling populations of rare or isolates species, wear disposable gloves and change them between handling each animal. Dedicate sets of nets, boots, traps, and other equipment to each site being visited. Clean and store them separately and the end of each field day.

5. When amphibians are collected, ensure the separation of animals from different sites and take great care to avoid indirect contact between them (e.g. via handling, reuse of containers) or with other captive animals. Isolation from un-sterilized plants or soils which have been taken from other sites is also essential. Always use disinfected/disposable husbandry equipment.

6. Examine collected amphibians for the presence of diseases and parasites soon after capture. Prior to their release or the release of any progeny, amphibians should be quarantined for a period and thoroughly screened for the presence of any potential disease agents.

7. Used cleaning materials (liquids, etc.) should be disposed of safely and if necessary taken back to the lab for proper disposal. Used disposable gloves should be retained for safe disposal in sealed bags.

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Source: Declining Amphibians Populations Task Force (DAPTF) http://www.amphibianark.org/pdf/Husbandry/The%20DAPTF%20Fieldwork%20Code%20of%20Practice.pdf

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Measuring Length

Salamanders: To measure the snout-vent length (SVL) of a terrestrial salamander (Fig. 2), hold it in your left hand (if you are right-handed) against your index and middle fingers with your thumb. This frees up the right hand for taking measurements. Hold the animal gently but firmly; if you hold it too tightly, it will struggle, but if you hold it too loosely, it will attempt to escape. When measuring the SVL, it is important that the vertebral column is straight. Measure from the tip of the snout to the anterior end of the vent to the nearest 0.1 mm with vernier calipers. If the anterior end of the vent correlates with the posterior insertion point of the hind legs, the latter may be easier to measure on the dorsal surface of the animal. Practice is needed to prefect this technique. If a salamander struggles, you may have to place it in a plastic bag to restrain it. Caution is needed when handling Ensatina because of their tendency to shed their tails when under stress (an energetically costly adaptation known as caudal autonomy).

Another method for restraining salamanders is the “Wall’s box.” The box consists of a plexiglas lid hinged to a box with a soft foam inside. The salamander can be “sandwiched” between the lid and the sponge. The salamander is thereby immobilized for measurement. As this method may be more stressful to the animal, and the foam bed maybe difficult to disinfect, it should only be used if accuracy is of high importance.

Fig. 2. Length Measurement (SVL) in Salamanders -ideally, the animal would be stretched out completely straight. Photo by K. Ovaska

Frogs: Standard measurements on frogs and toads are the head-body length (snout to posterior end of rump) and the head length (snout to posterior end of the tympanum), although other measurements might be taken depending on the purpose of the study. Measurements are usually taken to the nearest 0.1 mm with vernier calipers.

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Measuring Mass

Mass can be measured with a spring scale with the animal in a plastic bag, but a field electronic scale is usually more convenient and accurate. However, spring scales should be carried should the electronic scale fail or the animal is outside the range of the electronic scale. Mass is typically measured to the nearest 0.1g.

Releasing Amphibians

When releasing salamanders, place the animal on the ground and let it crawl under a cover object: don’t put the animal on the ground and put a cover object over it because the salamander might be pinned or crushed. Encourage the salamander to crawl under a cover object. This is important because salamanders left on the surface are subject to desiccation and predation.

When releasing anurans, just let them go at the point of capture – no special precautions are needed.

Marking Amphibians

The recognition of individuals or previously captured animals is essential for certain inventory objectives:

Estimating relative abundance through repeated sampling

Obtaining information on the movement of individual animals, growth rates, age at sexual maturity, frequency of reproduction and probability of survival

Monitoring population trends

For much inventory work, marking of animals is neither necessary nor desirable, as there are associated risks such as possible infection. Before undertaking a marking program, the reasons for doing so must be clearly considered.

Few amphibian species (e.g. Tiger Salamander) lend themselves to pattern mapping by possessing distinguishable, permanent, natural markings.

In the past, the most widely used method of marking amphibians and reptiles has been a form of mutilation known as toe clipping (Fig. 3). Although the potential detrimental effects have been well documented, the removal of one to eight digits is used extensively because it is inexpensive and quick. Detrimental effects can include pain, infection, and reduced agility. Toe clipping can reduce survival in the amphibians under study (McCarthy & Parris 2004).

Furthermore, toe clipping is unreliable because digits may regenerate (Donnelly et al. 1994), and larvae and the adults of certain species lack limbs. In a paper published in Nature, May (2004) questioned whether this practice is ethical.

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VANCOUVER ISLAND AMPHIBIANS

Source: Inventory Methods for Pond-breeding Amphibians and Painted Turtle (MELP, 1998)

Visible Implant Elastomer (VIE) is the most widely used alternative to toe clipping for identifying reptiles and amphibians. It is a two-part silicone-based material that is mixed immediately before use. VIE tags are injected as a liquid using a 28 gauge needle and cure into a pliable rubber-like material. Excess material is then wiped from the skin of the animal allowing the opening to heal and the tag to remain internal. Many researchers do not anesthetize amphibians, while others prefer the ease of handling that anesthetic provides, and still others have developed techniques in which the animal is placed in a plastic bag with some water and they tag through the bag.

Correct injection provides high retention and readability. Because the material is fluorescent it can be seen under pigmented skin and at night with the help of the VI Light. Anholt et al. (1998) state that the consistency and biocompatibility of the VIE tags allows for the tagging of small animals, including larvae, that could not be tagged using other methods. In a novel approach to monitoring the development of salamander egg masses, Regester and Woosley (2005) used VIE to identify and track the egg masses. Depending on temperature, the working time of the mixed elastomer is about 1-2 hours. While the elastomer will not be completely cured in that time, it will become very difficult to push it out of the injection syringe. The working time can be extended if the elastomer is stored in a freezer or on ice in a cooler after mixing.

VIE tags are an excellent choice for batch identification, and they also offer a surprising number of individual codes if colors and body locations are combined. For example, Jung et al. (2000) used three colors and four body locations to create 255 individual codes in his study of salamander. Successful tagging sites include between the toes in frogs, on the upper hind leg of frogs, and at the base of the limbs on the ventral side of salamanders. Lizards have been tagged in the upper legs and in the tail.

We will be practicing this technique on gummi frogs in Vertebrate lab, and you may be asked to devise a marking scheme for a population of salamanders during camp.

Figure 3. A symbolic coding scheme for toe-clipping.

The code is read from the left front foot to left back foot to right front foot to right back foot. The mark at the lower right reads 2403. The number of toes (usually 2 max/ individual), and which toes can be clipped, vary among species.

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Bibliography:Anholt, B.R., S. Negovetic, and C. Som. 1998. Methods for anaesthesia and marking of larval anurans.

Herpetological Review 29, 153-154

BC Wildlife Federation 2002. Conducting an Amphibian Inventory. Section five, Module 2.5. In The Wetlandkeepers Handbook. BC Wildlife Federation, Surrey, BC

Bury, R. B., and P. S. Corn. 1991. Sampling methods for amphibians in streams in the Pacific Northwest. General Technical Report PNW-GTR-275. USDA, Forest service, Pacific Northwest Research Station, Portland, Oregon.

Campbell, R. C. 1974. Statistics for Biologists. Cambridge University Press.

Corn, P. S., and R. B. Bury. 1990. Sampling methods for terrestrial amphibians and reptiles. General Technical Report PNW-GTR-256. USDA, Forest Service, Pacific Northwest Research Station, Portland, Oregon.

Declining Amphibians Populations Task Force (DAPTF) http://www.amphibianark.org/pdf/Husbandry/The%20DAPTF%20Fieldwork%20Code%20of%20Practice.pdf

DONNELLY, M. A., C. GUYER, J. E. JUTERBOCK, and R. A. ALFORD.1994. Techniques for marking amphibians. In W. R. Heyer, M. A. Donnelly, R. W. McDiarmid, L. C. Hayek, and M. S. Foster (eds.), Measuring and Monitoring Biological Diversity, Standard Methods for Amphibians, pp. 277-284. Smithsonian Inst. Press, Washington, D.C.

Heyer, W. R., M. A. Donnelly, R. W. McDiarmid, L.-A. C. Hayek and M. S. Foster. 1994. Measuring and monitoring biological diversity: standard methods for amphibians. Smithsonian Institution Press, Washington, D.C.

McCarthy, M.A. and K.M. Parris. 2004. Clarifying the effect of toe clipping on frogs with Bayesian statistics. .Journal of Applied Ecology 2004 41, 780–786 © 2004 British Ecological Society

Matsuda, B. M., D. M. Green and P. T. Gregory. 2006. Amphibians and Reptiles of British Columbia. Royal BC Museum, Victoria, BC.

MAY, R.M. 2004. Ethics and amphibians. Nature 431 :403.

MELP, 1998. Inventory Methods for Pond-breeding Amphibians and Painted Turtle. Standards for Components of British Columbia's Biodiversity No. 37. Prepared by Ministry of Environment, Lands and Parks Resources Inventory Branch for the Terrestrial Ecosystems Task Force March 13 1998 Version: 2.0

Olson, D. H., W. P. Leonard and R. B. Bury. 1997. Sampling amphibians in lentic habitats: methods and approaches for the Pacific Northwest. Society for Northwestern Vertebrate Biology, Olympia, Washington.

Ovaska, Kristiina: http://saltspringconservancy.ca/wp/plants-animals/salt-spring-wildlife/amphibians (Fig. 2 photo credit)

Regester, Kurt J., Woosley, Lori B. 2005. Marking Salamander Egg Masses with Visible Fluorescent Elastomer: Retention Time and Effect on Embryonic Development. The American Midland Naturalist 153(1):52-60

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Further ReadingDavis, T. 1997a. Standardized inventory methodologies for components of British Columbia's

biodiversity: terrestrial salamanders. Ministry of Environment, Lands and Parks, Resources Inventory Branch, Victoria, B.C.

Davis, T. M. 1997b. Non-disruptive monitoring of terrestrial salamanders with artificial cover objects on southern Vancouver Island, British Columbia. In D. M. Green (ed.), Amphibians in decline: Canadian studies of a global problem. Herpetological Conservation 1:161-174.

Davis, T. M. 1999. Study designs for evaluating the effects of forestry activities on aquatic-breeding amphibians in terrestrial forest habitats of British Columbia. Ministry of Environment, Lands and Parks, Wildlife Branch, Victoria, B.C. 42pp.

Davis, T. M. 2002. Research Priorities for the Management of the Western Toad, Bufo boreas, in British Columbia. B.C. Minist. Water, Land and Air Protection, Biodiversity Branch, Victoria, BC. Wildlife Working Report No. WR-106.

Davis, T. M. and K. Ovaska. 2001. Individual recognition of amphibians: effects of toe clipping and fluorescent tagging on the salamander Plethodon vehiculum. Journal of Herpetology 35:217-225.

Fellers, G. M. and K. L. Freel. 1995. A standardized protocol for surveying aquatic amphibians. Technical Report No. NPS/WRUC/NRTR 95-01. United States Department of the Interior, National Park Service, National Biological Service, University of California.

Jones, L. L. C., W. P. Leonard and D. H. Olson. 2005. Amphibians of the Pacific Northwest. Seattle Audubon Society, Seattle, USA.

Stebbins, R.C. and N.W. Cohen. 1995. A Natural History of Amphibians. Princeton University Press, New Jersey.

Amphibians of Vancouver Island and the Gulf Islands: http://www.saltspringconservancy.ca/pdf/Amphibian%20brochure,%20revised%20Aug%2012.pdf

BC Species and Ecosystems Explorer: http://www.env.gov.bc.ca/atrisk/toolintro.html

Canadian Herpetological Society: http://www.carcnet.ca/english/amphibians/tour/province/amphBC.php

E-Fauna BC: http://www.geog.ubc.ca/biodiversity/efauna/amphibians.html

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