Wetland Wildlife Monitoring and Assessment

Abstract

Monitoring wetland wildlife is complex and requires use of various techniques to obtain robust population estimates. Herpetofauna, birds and mammals frequently inhabit wetlands and adjacent uplands. Sampling herpetofauna can include passive techniques such as visual encounter and breeding call surveys, and capture techniques that use nets and traps. Common bird monitoring techniques include scan surveys, point counts, nest searches, and aerial surveys. Some mammals, such as bats, can be sampled with audio devices, whereas mark-recapture techniques are most effective for other taxa. For all groups, the techniques used depend on the monitoring objective and target species. This chapter describes various techniques for monitoring populations of wetland wildlife. If these techniques are incorporated into a robust sampling design, they can be used to document changes in species occurrence, relative abundance, and survival.

Student Exercises

7.1.1 Laboratory Exercise #1: Herpetofaunal Sampling Laboratory

Location and Time: Herbaceous or forested wetland with standing water during the growing season (ideally spring or summer).

Description: The goal of this field lab will be to build competency in common techniques used to sample amphibian and reptile communities. Activities described herein could be assigned in their entirety or only portions of the exercise used. If completed in its entirety, students will have an understanding how to use pitfalls, funnel traps, and nets to capture amphibians, use basking traps to capture turtles, and collect biological information on captured amphibians and turtles.

Supplies and Equipment: Silt fencing with attached wooden stakes, 3-lb sledge hammer, rake, tape measure, shovel, six 19-L (5-gal) buckets, six sponges, six rectangular funnel traps, dip nets, seine net, basking turtle trap, snake tongs, calipers, organism bags, spring scales, and disposable gloves (worn while handling animals). Given that live animals will be captured and handled, scientific collection permits must be secured, and an Institutional Animal Care and Use Committee (IACUC) protocol may be required by your institution.

Set-up Instructions: Identify a wetland for sampling and remove leaves, debris, and herbaceous vegetation in a 0.5-m wide band 10 m above the high waterline and parallel to the wetland for 40 m. Dig holes for 5-gal pitfalls every 10 m, with two pitfalls paired at each end. Pitfall tops should be flush with the top of the ground. Erect fencing such that it passes next to each pitfall and 0.5 m past the end pitfalls. Cover the base of the fence with soil to prevent trespass of animals. Fill pitfalls with 2.5 cm of water from the wetland and place one sponge in each pitfall. Place one funnel trap on each side of the fence, and one trap in shallow water (<10 cm) in each cardinal quadrant of the wetland. Construct a basking trap following Brown and Hecnar (2005), and place in water >1 m depth.

Sampling Instructions: After deployment of traps, check in <24 h. Identify all captured juvenile and adult amphibians and reptiles in pitfall and funnel traps, and measure the snout-to-vent length and mass. Amphibians can be placed in plastic bags when processing but should be rehydrated with water from the wetland before release. Lizards can be placed in cloth bags or plastic containers. When handling lizards, avoid grabbing by the tail because most species will autotomize it as an anti-predator response. Captured snakes should only be handled after verifying they are not venomous; non-venomous snakes can be temporarily held in a well-secured pillowcase. Venomous snakes should not be handled and be removed from traps using snake tongs or snake tubes. Aquatic and terrestrial turtles can be placed in 5-gal buckets or large plastic containers. Care should be taken when handling snapping turtles because their bite can cause injury. Larval amphibians can be sampled using dip and seine nets following Schmutzer et al. (2008). Identify and enumerate all larval amphibians by species. As a second exercise, determine developmental stage according to Gosner (1960).

Brown CL, Hecnar SJ (2005) Capture success of northern map turtles (Graptemys geographica) and other turtle species in basking vs. baited hoop traps. Herpetol Rev 36:145–147

Gosner, KL (1960) A simplified table for staging Anuran embryos and larvae with notes on identification. Herpetologica 16:183–190

Schmutzer AC, Gray MJ, Burton EC, Miller DL (2008) Impacts of cattle on amphibian larvae and the aquatic environment. Freshw Biol 53:2613–2625

7.1.2 Laboratory Exercise #2: Amphibian Marking Laboratory

Location and Time: This laboratory can be completed in combination with the Herpetofaunal Sampling Laboratory or in the classroom with preserved laboratory specimens.

Description: The purpose of this laboratory is to practice common herpetofaunal marking methods. Activities in this laboratory can include marking techniques for adult or larval anurans. If this laboratory is completed in its entirety, students will have the experience necessary to mark amphibians using scissors, Visible Implant Elastomers (VIE), and Passive Integrated Transponder (PIT) tags.

Supplies: Wild captured or preserved amphibians (3 per student or team), surgical grade scissors, VIE mix and injection syringes (Northwest Marine Technologies, Inc.), PIT tag supplies (PIT tags, tag reader, and injection syringe [Biomark, Inc.]), 95 % EtOH or 2 % chlorhexidine diacetate solution, disposable gloves, and appropriate marking schemes. If live amphibians are captured, scientific collection permits must be secured, and an IACUC protocol may be required by your institution.

Set-up and Instructions: Wild amphibians can be captured during the Herpetofaunal Sampling Laboratory, and preserved specimens (e.g., Necturus, Lithobates) can be acquired from most biological supply companies. If wild animals are used, it is important to sanitize all marking instruments using 95 % ETOH or 2 % chlorhexidine diacetate solution. Prior to the lab, review and select one of the published marking schemes for amphibians (see Donnelly et al. 1994 or Ferner 2010). Students should work on a stable, clean surface. Distribute at least three amphibians to each student or group (i.e., one amphibian per marking technique). Students should wear disposable gloves and change them between animals. Practice toe clipping as described in the chapter. The excision should be made at the most proximal joint; bones and thumbs should not be cut. Excise the appropriate toes to number the individual as #234. Once completed, use a different animal for VIE marking. The elastomer should be injected under the skin where very little pigment occurs; the ventral side of most amphibian legs is a good location. Care should be taken to slide the needle under the skin (forceps can help) so as to not pierce muscle or organs. When working with live amphibians, students should work in pairs, with one student holding the animal securely. Students should conceive schemes that allow for individual or batch marking using different VIE colors and marking locations on the amphibian. Finally, practice injecting a PIT tag under the skin mid-body on ventral and dorsal sides. Scan the tag prior to and after injecting. At the conclusion of the laboratory, students should discuss advantages and disadvantages of each technique. Processing time for each technique should be recorded and considered.

Donnelly MA, Guyer C, Juterbock JE, Alford RA (1994) Techniques for marking amphibians. In: Heyer WR, Donnelly MA, McDiarmid RW, Hayek LAC, Foster MS (eds) Measuring and monitoring biological diversity: standard methods for amphibians. Smithsonian Institution Press. Washington, DC, pp 277–284

Ferner JW (2010) Measuring and marking post-metamorphic amphibians. In: Dodd CK (ed) Amphibian ecology and conservation: a handbook of techniques. Oxford University Press, Oxford, pp 123–142

7.1.3 Laboratory Exercise #3: Small Mammal Trapping Laboratory

Location and Time: Perimeter of a forested or emergent wetland in areas not subject to permanent flooding, or the adjacent upland can be used; performed preferably during fall.

Description: The goal of this lab will be to expose students to basic live-trapping techniques used to estimate abundance, density, or distribution of small mammals. This lab will require students to establish trapping grids prior to setting traps, set traps in the afternoon, and check all traps the following morning. Students will gain an understanding of how to establish trapping grids, set traps, capture and mark small mammals, and collect basic morphological and demographic data.

Supplies: Sherman or Longworth traps, flagging tape, peanut butter and oats, wax paper, clear plastic bags, spring scale, and disposable gloves. Given that live animals will be captured and handled, scientific collection permits must be secured, and an IACUC protocol may be required by your institution.

Set-up Instructions: Identify an area that is not permanently flooded with suitable herbaceous or woody vegetation to harbor small mammals. Establish a trapping grid with a minimum of 25 traps (e.g., 5 × 5 matrix) with each trap placed 10 m apart. Place flagging at each trap site so that traps can be quickly relocated. Prior to setup, mix oats and peanut butter together. Cut 3 × 3 in. squares of wax paper, and place ½ teaspoon of peanut butter mixture in center of wax paper, fold the ends together, and twist so that the peanut butter mixture is inside a pouch of wax paper.

Bait each trap by placing the bait balls (inside of wax paper) at the back of each trap. Traps should be placed in areas likely to be used by small mammals, such downed woody debris, stumps, etc. Place one trap at each trapping site during the afternoon or evening prior to the day that traps will be checked. Trap sites should be spatially referenced using a GPS unit to ensure re-location of the sites during monitoring.

Sampling Instructions: All traps should be checked <16 h after being set. Mammals captured in traps can be removed by opening the door of the trap, placing a clear plastic bag over the trap door, and turning the trap over to drop the captured individual into the bag. Each person handling mammals should wear disposable gloves. Mammals can be carefully removed from the bag by pinching the fur around the back of the neck. Larger mammals, such as cotton rats (Sigmodon hispidus), can be further restrained by holding their tails with the other hand. Identify mammals to species and determine sex. Small mammals can be identified using Peterson (2006). Body weight can be determined by closing the plastic bag briefly, and hanging the bag from a spring scale. Capture data should be summarized by species, and inferences made about their association with different cover types. For instance, students could simultaneously measure habitat characteristics (e.g., canopy cover, vegetation type, vegetation density) at sites where mammals are successfully captured and compare them to sites where mammals are not captured.

Peterson RT (2006) Peterson field guide to mammals of North America, 4th edn. Houghton-Mifflin, Boston

7.1.4 Laboratory Exercise #4: Evaluating Wildlife Sign for Surveys

Location and Time: In-class laboratory with slide presentation.

Description: The goal of this lab is to train students to identify tracks and sign of common mammals likely to be encountered in various wetland habitats. The slide presentation is designed to provide students with information on how to identify tracks based on numbers of toes, distance between front and rear feet, morphological characteristics of feet among species, as well as gait patterns. The lab is most effective if students have plaster casts of the species covered in the presentation so that they can view tracks and study them.

Supplies: Slide presentation (PDF format) by Mike Chamberlain is available for use at: http://fwf.ag.utk.edu/mgray/WetlandBook/WildlifeSignsLab.pdf. If track casts are unavailable, a reference collection can be created using Plaster of Paris available at craft stores.

Set-up Instructions: The presentation describes how to identify tracks of mammals based on numbers of toes. Specifically, students should be instructed on ways to recognize 2-toed hooved species, 4-toed species with heal pads, 5-fingered species, and species with 4 front toes and 5 hind toes. Mammals occupying wetlands vary by locale, but larger, more common species, such as white-tailed deer, feral hog, coyote, red fox, gray fox, bobcat, cottontail and swamp rabbits, raccoon, opossum, muskrat, beaver, mink, river otter, and black bear, are covered in this lab.

7.1.5 Laboratory Exercise #5: Waterbird Identification, Sexing and Aging Laboratory

Location and Time: Indoor laboratory during fall or spring. This lab should be conducted prior to the Waterbird Population Monitoring field lab.

Description: The goal of this lab will be to expose students to waterbirds that are of management and conservation interest in the state and region. In addition, sexing and aging techniques by plumage will be demonstrated for species where this information is of management interest, such as waterfowl. The students will be responsible for identification of the species that are presented in the lab and sexing and aging for a subset of those species.

Supplies: Photographs of species of interest and study skins where possible, waterfowl wings for males and females of species of interest, bird field guide, and waterfowl wing sexing and aging guide (Carney 1992). Also, a slide presentation by Matthew Gray on the identification of North American waterfowl is available: http://fwf.ag.utk.edu/mgray/wfs560/WaterfowlID.pdf.

Classroom Instruction: Develop a list of waterbirds that students will be responsible for learning including waterfowl, wading birds, shorebirds, and secretive marsh birds. Include species that are generally of management interest for the state or region, including species that will likely be encountered during the field lab. Develop a slide presentation in which the instructor reviews the identification characteristics of the species on the list. The instructor should also review the sexing and aging techniques for species of interest. After the presentation is complete, the students will break into 2-person teams to review the specimens that are available using their field guides to make a positive identification. In addition, they can use the U.S. Fish and Wildlife Service sexing and aging guide for waterfowl wings as additional practice (see below).

Lab Proficiency Quiz: When each student (or team) believes they have mastered identification, they can attempt a proficiency quiz. The quiz should include images of birds at varying distances. They will be declared proficient if they correctly identify >70 % of the birds. Students should demonstrate proficiency prior to the field lab.

Carney SM (1992) Species, age and sex identification of ducks using wing plumage. U. S. Department of the Interior, U.S. Fish and Wildlife Service, Washington, DC. Northern Prairie Wildlife Research Center, Jamestown. http://www.npwrc.usgs.gov/resource/tools/duckplum/index.htm

7.1.6 Laboratory Exercise #6: Waterbird Population Survey Laboratory

Location and Time: Wetlands with open water and mudflats during fall or spring.

Description: The goal of this field lab will be to demonstrate population survey methods for waterfowl, shorebirds and wading birds, and allow students to practice the methods, analyze the data, and interpret the results.

Supplies: Binoculars and spotting scopes, laser rangefinders, study area maps, bird field guides, clipboards, data sheets, and 1-m stakes for each student team.

Set-up Instructions: Identify a wetland with open water and mudflats for survey. Divide the wetland area into sampling units based on geographic area (e.g., cardinal quadrants) or habitat. Use the maps, laser rangefinders, and stakes to delineate the spatial extent of each survey location. Each location should survey approximately the same viewable area. Divide the class into 2–4 person teams and assign each team to a location.

Survey Instructions: At the beginning of the laboratory, explain the goals of the exercise and review the count protocols and identification for species likely to be encountered. If there are species that are difficult for novices to identify (e.g., various sandpiper species), group these as morpho-species (e.g., western, least and semi-palmated sandpipers might be counted simply as “sandpipers”). Deploy the teams to conduct the counts, ideally within 3 h of sunrise or sunset. Each team should spend the first 30 min scanning the area and identifying the waterfowl, wading birds and shorebirds to species. The last 15 min of the count period will be spent estimating a count for each species. As an additional exercise that students can practice focal surveys, where bird activities are recorded (e.g., feeding, walking, swimming, inactive, sleeping, antagonistic, alert) for 1 min. Students are encouraged to read Davis and Smith (1998) for an example of collecting and analyzing activity budget data.

Data Analyses: Data from each team should be entered into a database, including date, time, study area(s), environmental conditions (temperature, wind speed and direction, precent cloud cover, precipitation), observers, species and counts. Each team will summarize the data collected by the entire class to make inferences about waterbird use of the area. After conducting this exercise over several years, students can begin to look for seasonal or yearly changes in species composition and abundance, and develop hypotheses for why these demographics may be fluctuating.

Written Assignment: Each team will be responsible for writing up a lab report, summarizing the objectives, methods, and results from the surveys and discussing the implications of the results and answering critical questions about changes in waterbird populations. Additionally, each team may present their results orally, and a class discussion can explore the lessons learned from this experience.

Davis CA, Smith LM (1998) Behavior of migrant shorebirds in playas of the Southern High Plains. Condor 100:266–276