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https://www.proquest.com/docview/2314099870?accountid=15112&parentSessionId=PmD%2B4rkn4Y3%2BPG0aA4m0L%2FruBLmZzBTAQaZPiDZnf30%3D&pq-origsite=primo

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https://www-sciencedirect-com.ezproxy.uow.edu.au/science/article/pii/S1470160X1630526X?via%3Dihub

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fp_cmer_04_402.pdf (wa.gov)

Comparison of Three Methods for Surveying Amphibians in Forested Seep Habitats in Washington

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Study looks at the assemblage composition, seasonal abundance and vertical distribution of litter-dwelling thrips.

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Surveying techniques arboreal vs grounddwellign mammals

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Ecology, management and conservation in natural and modified habitats

P. C. Catling, R. J. Burt and R. Kooyman

Wildlife Research 24(4) 417 - 432
Published: 1997

Abstract

An extensive fauna survey of forest habitats in the Murwillumbah Forestry Management Area of north-eastern New South Wales allowed us to examine and compare the use of Elliott traps, wire cage traps, soil plots, hair tubes, spotlighting, dry pitfall traps, sightings and vocalisations as methods to determine the identity, distribution and abundance of ground-dwelling and arboreal mammals. Transects were established in 10 areas in six State Forests and one National Park.

The aim of this study was to examine each method to satisfy three criteria and to determine the most efficient and effective method or combination of methods to survey arboreal and ground-dwelling mammals in forests. The three criteria were to identify the species present, to determine their distribution and to establish an index of their abundance. In all, 35 species were identified by the seven methods. However, the species recorded, and the usefulness of the method to provide their distribution and/or an index of abundance, differed markedly according to the method used.

Stratified sampling with Elliott traps for small mammals, spotlights for arboreal mammals and soil plots for medium-sized and large ground-dwelling mammals was the most labour-efficient and productive combination for recording species present and their status comprehensively. If needed, other methods such as hair tubes or cage traps may be used locally and intensively to assist in the identification of species. Moreover, randomly gathered sight and vocalisation records accumulated during the survey were found to be very useful in corroborating species present and in helping to identify the tracks recorded to genus or family by the soil-plot method.

https://doi.org/10.1071/WR96073

© CSIRO 1997

CSIRO Wildlife Research

All volumes of CSIRO Wildlife Research are online and available to subscribers of Wildlife Research.

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A Comparison of Techniques Used in a Survey of the Ground-dwelling and Arboreal Mammals in Forests in North-eastern New South Wales

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An extensive fauna survey of forest habitats in the Murwillumbah Forestry Management Area of north-eastern New South Wales allowed us to examine and compare the use of Elliott traps, wire cage traps, soil plots, hair tubes, spotlighting, dry pitfall traps, sightings and vocalisations as methods to determine the identity, distribution and abundance of ground-dwelling and arboreal mammals.

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Bat Surveying Guidelines Australia + Additional Information Regarding Taxon

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Bat Surveying Guidelines Australia + Additional Information Regarding Taxon

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Survey guidelines for Australia's threatened bats (agriculture.gov.au)
As bats may prove a difficult species to catch for surveying purposes, the range and extremity of surveying methods is broad. Including, trip lines, mistnets and roost searches. Document details various methods used, aswell as the steps to complete successful surveying.

Selection of roost sites by the lesser long‐eared bat (Nyctophilus geoffroyi) and Gould's wattled bat (Chalinolobus gouldii) in south‐eastern Australia (wiley.com)
Information relating to the roosting habits of bats could prove useful in a surveying attempt that relied upon roost searches.

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General Bird Survey - USGS Breeding Bird Surveys

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General Bird Survey - USGS Breeding Bird Surveys

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Similar to simple transects, this survey method involves travelling on North American roads over a pre-determined distance (24.5 miles) and stopping each 0.5 mile for 3 minutes to measure bird abundance and diversity.

https://www.pwrc.usgs.gov/bbs/about/

Primary literature which integrates this survey method - https://doi-org.ezproxy.uow.edu.au/10.1016/j.foreco.2007.02.016

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About BBS

What is the North American Breeding Bird Survey (BBS)?
The BBS is a long-term, large-scale, international avian monitoring program initiated in 1966 to track the status and trends of North American bird populations. The USGSPatuxent Wildlife Research Center and the Canadian Wildlife Service, National Wildlife Research Center jointly coordinate the BBS program.
Why was the BBS created?
In the mid-twentieth century, the success of DDT as a pesticide ushered in a new era of synthetic chemical pest control. As pesticide use grew, concerns, as epitomized by Rachel Carson in Silent Spring, regarding their effects on wildlife began to surface. Local studies had attributed some bird kills to pesticides, but it was unclear how, or if, bird populations were being affected at regional or national levels. Responding to this concern, Chandler Robbins and colleagues at the Patuxent Wildlife Research Center developed the North American Breeding Bird Survey to monitor bird populations over large geographic areas.

Although most concerns over pesticide use in North America have subsided in recent decades, bird populations continue to be subjected to numerous widespread threats including habitat loss, habitat fragmentation, land-use changes, and other chemical contaminants. Today, the BBS continues to monitor bird populations across North America and informs researchers and wildlife managers of significant changes in bird population levels. If significant declines are detected, their causes can then be identified and appropriate actions taken to reverse them before populations reach critically low levels.

How does the BBS work?
Each year during the height of the avian breeding season, June for most of the U.S. and Canada, participants skilled in avian identification collect bird population data along roadside survey routes. Each survey route is approximately 24.5 miles long with stops situated ideally 0.5-mile apart. At each stop, a 3-minute point count is conducted. During the count, every bird seen within a 0.25-mile radius or heard is recorded. Surveys start one-half hour before local sunrise and take about 5 hours to complete. Over 4100 survey routes are located across the continental U.S. and Canada.

Once analyzed, BBS data provide an index of population abundance that can be used to estimate population trends and relative abundances at various geographic scales. Trend estimates for more than 420 bird species and all raw data are currently available via the BBS web site.
How are BBS data used?

1. The U.S. Fish and Wildlife Service,Canadian Wildlife Service, and Partners in Flight all use BBS trends along with other indicators to assess bird conservation priorities.

2. BBS data were instrumental in focusing research and management action on neotropical migrant species in the late 1980s, and on grassland species in the mid-1990s.

3. State Natural Heritage programs and Breeding Bird Atlas projects often utilize BBS data to enrich their databases.

4. Educators often use BBS data as a tool to teach biological, statistical and GIS concepts.

5. More than 450 scientific publications have relied heavily, if not entirely, on BBS data. The entire BBS bibliography is viewable here.

|
Laurel, MD, USA 20708-4038
http://www.pwrc.usgs.gov/bbs
Contact Us
Last Modified: 3/27/2018
BBS Privacy Statement

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BBS - About - USGS Patuxent Wildlife Research Center

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About BBS What is the North American Breeding Bird Survey (BBS)? The BBS is a long-term, large-scale, international avian monitoring program initiated in 1966 to track the status and trends of North American bird populations. The USGSPatuxent Wildlife Research Center and the Canadian Wildlife Service, National Wildlife Research Center jointly coordinate the BBS program.Why was the BBS created?

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Threatened species survey and assessment guidelines: field survey methods for fauna - Amphibians | NSW Environment and Heritage

These are specific guide-lines for surveying endangered amphibian species. Different species show different seasonal or daily peaks in activity and this is something really important to take into account whilst surveying.

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Survey Guideline
https://www.dcceew.gov.au/sites/default/files/documents/survey-guidelines-birds-april-2017.pdf

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Community structure, species-habitat relationships, and conservation of amphibians in forested vernal pools

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Community structure, species-habitat relationships, and conservation of amphibians in forested vernal pools

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Study involves survey on abundance of breeding amphibians, investigating effects of habitat qualities (hydroperiod length, canopy openness, environmental vegetation, etc.) on abundance.

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https://www.frontiersin.org/articles/10.3389/fevo.2020.579193/full
Summarizes best practice for surveying insects

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Department of Ecology and Evolutionary Biology, University of California, Los Angeles, Los Angeles, CA, United States

Florida Museum of Natural History, University of Florida, Gainesville, FL, United States

Department of Biology, University of Florida, Gainesville, FL, United States

Benchmark studies of insect populations are increasingly relevant and needed amid accelerating concern about insect trends in the Anthropocene. The growing recognition that insect populations may be in decline has given rise to a renewed call for insect population monitoring by scientists, and a desire from the broader public to participate in insect surveys. However, due to the immense diversity of insects and a vast assortment of data collection methods, there is a general lack of standardization in insect monitoring methods, such that a sudden and unplanned expansion of data collection may fail to meet its ecological potential or conservation needs without a coordinated focus on standards and best practices. To begin to address this problem, we provide simple guidelines for maximizing return on proven inventory methods that will provide insect benchmarking data suitable for a variety of ecological responses, including occurrence and distribution, phenology, abundance and biomass, and diversity and species composition. To track these responses, we present seven primary insect sampling methods-malaise trapping, light trapping, pan trapping, pitfall trappings, beating sheets, acoustic monitoring, and active visual surveys-and recommend standards while highlighting examples of model programs. For each method, we discuss key topics such as recommended spatial and temporal scales of sampling, important metadata to track, and degree of replication needed to produce rigorous estimates of ecological responses. We additionally suggest protocols for scalable insect monitoring, from backyards to national parks. Overall, we aim to compile a resource that can be used by diverse individuals and organizations seeking to initiate or improve insect monitoring programs in this era of rapid change.

Introduction

" The best time to plant a tree is 20 years ago. The second best time is now." -Unattributed proverb

The threat of widespread insect declines, supported by accumulating evidence across the globe ( Conrad et al., 2006; Forister et al., 2011; Hallmann et al., 2017; van Klink et al., 2020), has sparked broad and outspoken concern. But even this general pattern of insect decline is heterogeneous in time and space, and drivers of declines in particular taxa and locations remain unclear, though they are likely myriad ( Fox, 2013; Wagner, 2020). To better understand insect declines in the face of data gaps and other challenges ( Didham et al., 2020), researchers need more systematic and long-term monitoring of insect abundance and diversity. Though many monitoring schemes already exist, relatively few have been operating long enough to draw robust, independent conclusions about insect populations and diversity over time (e.g., Shortall et al., 2009), and these monitoring schemes are necessarily limited in their geographic and taxonomic coverage. As scientists, we can and should lament our severely limited data on insect declines-long-term monitoring efforts should have been underway long before now, but were prevented for many reasons, including a lack of funding, motivation, and organization. Given increased societal interest in insects, there is the potential for widespread, long-term monitoring at the scale necessary to benchmark and track insect trends moving forward.

Many new monitoring efforts have been recently initiated, motivated by reports of insect declines. Researchers, managers, and community scientists are currently increasing efforts to document insects, whether photographing insects at porch lights, counting pollinators on transects, or establishing structured malaise trap programs. These efforts are a crucial first step toward broadly tracking trends in insect abundance and diversity. To maximize the information gain from these largely independent efforts, we recommend integration with established insect monitoring methods to coordinate sharing data that are accessible and interoperable. As much as possible, new monitoring efforts should align methods, metadata, and data access with those that already exist to increase explanatory power, streamline analysis, and facilitate the development of a global insect monitoring network. This network is already beginning to form through the efforts of organizations like PollardBase ( Taron and Ries, 2015), the National Moth Recording Scheme ( Fox et al., 2011), the Global Malaise Program ( Geiger et al., 2016), as well as regional efforts, taxon-specific programs (e.g., for monarch butterflies and lady beetles), and even groups of Twitter users organizing nights to check their porch lights for insects. Recently, Woodard et al. (2020) took the important step of proposing a national bee monitoring network in the United States. However, the urgency of insect declines requires even more rapid development and integration in an era of purported "insect apocalypse."

To reach the goal of a long-term monitoring network on a global scale, we will need data that, through standardization and well-defined metadata, can be integrated across monitoring efforts. Without standardized data and metadata collection, researchers will assemble datasets that are difficult or impossible to integrate, hindering synthesis. In other words, the efforts of thousands working independently are most valuable when those efforts can be assembled into a collective whole.

To meet this challenge, our aim here is to inform new monitoring projects with standardized data collection and metadata collection practices, facilitating future integration. We present a standardized toolbox for monitoring methods and metadata practices, aimed as a starting point for non-specialists and a reference point for specialists. Specifically, we provide: (1) overviews of common insect monitoring methods, including malaise, light, pan, and pitfall trapping, beating sheet, and audio and active visual surveys; (2) specific recommendations for how to carry out each method in the field; (3) an overview of metadata considerations; (4) recommendations for standardized metadata collection for each method ( Table 1); and (5) a forecast of emerging methods that can complement and extend existing methods.

Table 1. Recommended metadata for each of seven methods of insect monitoring.

Our audience is anyone interested in insect monitoring, from community members motivated to contribute to science, to entomological specialists who want to make their data more broadly useful. We especially hope these recommendations will aid those interested in insect monitoring but are not sure where to start. Workers can choose monitoring methods from the toolbox we present, then modify as needed for their goals and systems. We make these recommendations with scalability in mind-the methods we discuss are generally low cost, field-tested, and can be performed by a single individual. We generally organize these methods by following the framework presented by Ferro and Summerlin (2019), while our summaries and recommendations are especially influenced by Southwood and Henderson (2000), Samways et al. (2010), and previous efforts to advocate for sampling alignment for bees ( Droege et al., 2017) and birds ( Ralph et al., 1993). Along these lines, we do not advocate for existing monitoring networks to change their methods even if they are not easy to integrate with other efforts. Though methodological standardization is ultimately a goal, we pragmatically advocate as much for alignment with and among existing monitoring efforts as we do for standardization.

In the following sections, we outline recommended methods for sampling a given set of insect taxa for monitoring purposes. These methods are generally suitable for a variety of key benchmarking goals, including the measurement of occurrence and distribution, phenology, abundance and biomass, and diversity and species composition ( Box 1). This is not a guide for conducting exhaustive species inventories (e.g., BioBlitzes or site lists), which often emphasize maximizing species counts, nor is it a guide for maximizing insect catches. This guide also does not aim to eliminate bias-no method is free of bias, but if methods and metadata are documented carefully and are consistent over time, then bias can largely be estimated and controlled. Additionally, monitoring insects in sites with rare and endangered species also requires unique considerations that must be site- and species-specific, and we do not cover the complexities of those considerations here, nor do we go into details of how to analyze data once collected. Finally, we do not recommend these sampling methods for entomologists with highly specific taxonomic goals; the class Insecta is simply too diverse in its niches and behaviors to be comprehensively assessed by one or even a few sampling methods. Here, we attempt to create a balance between being too general or too specific, by presenting a variety of methods for sampling broad taxa and guilds, each with its own strengths and weaknesses ( Figure 1). If you find yourself wanting to monitor more specific insect groups, your sampling methods may need to be modified from those discussed here or may not be covered.

Box 1. Measurement goals of benchmarking studies.

The goals of benchmarking and monitoring studies typically aim to measure change in at least one of the following: (1) occurrence and distribution, (2) phenology, (3) abundance and biomass, or (4) diversity and species composition. The seven monitoring methods we highlight can be used with the goal of measuring changes in as many of these responses as desired. Each of these four categories of response is important for different reasons, requires a different minimum spatiotemporal scale of sampling, and is currently used to study how insect populations and communities change over time.

Occurrence and distribution: Changes in occurrence and distribution are important indicators of how shifts in underlying processes affect organisms. Occurrence can also sometimes serve as proxies for abundance ( Royle and Nichols, 2003). Estimating occurrence and distribution requires, at minimum, the formal identification of a taxon at a location (i.e., a presence), but the addition of data on what taxa were not present (i.e., an absence), allows for a more powerful analysis of occurrence. For insects, occurrence and distribution monitoring is perhaps the most widespread benchmarking method (e.g., Chen et al., 2009; Boyes et al., 2019; Outhwaite et al., 2019), especially for invasive species and in the context of shifting ranges due to climate change.

Phenology: Shifts in phenology can indicate changes in the factors governing the timing of insect life cycles, from temperature and precipitation patterns, to flowering periods in plants. Shifts in insect phenology can cause mismatches with other taxa in their communities, from plants to birds, and can have demographic consequences for those taxa ( Visser and Gienapp, 2019). Estimating phenology typically requires, at minimum, presence data for a taxon at a location repeatedly over a short time span (i.e., a "season"), but presence and absence data together allow for stronger inference. Changes in insect phenology are poorly documented in most taxa, but recent interest in the effects of climate change has spurred a larger focus on insect phenology in monitoring efforts ( Gimesi, 2012).

Abundance and biomass: Changes in abundance and biomass, both measures of ecosystem function, are important for understanding the health of the ecosystem as well as for conservation and management. Estimating abundance and biomass typically requires presence and absence data in addition to accurate counts of individuals of each taxa. This form of monitoring is historically rare but has been perhaps the most influential in spurring recent interest in insect declines ( Hallmann et al., 2017; Wepprich et al., 2019).

Diversity and species composition: Changes in measures of biodiversity, such as species diversity and composition, indicate how communities respond to environmental change. Estimating diversity and species composition typically requires presence and absence data for multiple taxa in a community, with some expectation that sampling is equally likely across taxa. Monitoring insect diversity and composition is relatively common compared to abundance, biomass, and phenological monitoring (e.g., Brooks et al., 2012; Valtonen et al., 2017).

Figure 1. Commonly-monitored insect guilds and taxa and the efficacy for each of seven benchmarking methods. Efficacy of each method for a given insect group is scored as follows: filled green circles indicate optimal suitability; half-filled circles indicate possible suitability; divided, unfilled circles indicate marginal suitability; unfilled gray circles indicate bycatch only; and no circle indicates general unsuitability. Insect groupings are defined by ecological traits (blue bar) or taxonomic clades (purple bar). In order by column, insect groupings are: adult semi-aquatic insects (Plecoptera, Ephemeroptera, and Trichoptera); singing insects (Orthoptera & Hemiptera: Cicadoidea); ground-dwelling beetles (Coleoptera: Carabidae and Staphylinidae); non-lepidopteran pollinators (Hymenoptera, Diptera, Coleoptera); leaf-chewing larvae (Lepidoptera and Hymenoptera: Symphyta); night-active moths (Lepidoptera); dragonflies and damselflies (Odonata); ants (Hymenoptera: Formicidae); butterflies (Lepidoptera: Papilionoidea); and flies (Diptera).

Standardized Monitoring Practices for Different Taxa and Methods

Malaise Trapping

Overview

Malaise traps ( Malaise, 1937) are large tent-like structures made of netting meant to funnel insects to a common area ( Figure 2A). In essence, an insect flies into a vertical wall of netting, responds by flying upwards, then is gradually funneled by sloped netting into a collecting vial. This vial is then checked and emptied periodically over days or weeks. The Townes-type malaise trap is the most common style used, but at least four other types-Gressitt malaise, Schact malaise, Sea, land, and air malaise (SLAM), and Cornell malaise-are also in use ( Matthews and Matthews, 1983; van Achterberg, 2009). For detailed accounts of history and methodology, see van Achterberg (2009). Exemplar malaise trapping programs include the School Malaise Trap Program in Canada ( Steinke et al., 2017) and the Swedish Malaise Trap Program ( Karlsson et al., 2020). For those interested in joining an existing network, the Global Malaise Trap Program/BIOSCAN ( Geiger et al., 2016) is accepting new members.

Figure 2. A visual overview of the seven insect benchmarking methods summarized here: (A) malaise trapping, (B) light trapping, (C) pan trapping, (D) pitfall trapping, (E) beating sheet, (F) acoustic monitoring, and (G) active visual surveys.

Taxonomic Considerations

Malaise trapping is only appropriate for monitoring flying insects ( Figure 1). Many flies (Diptera) and some wasps, flying ants, bees (Hymenoptera), bugs (Hemiptera), moths (Lepidoptera), and semi-aquatic taxa are effectively sampled by malaise traps ( Matthews and Matthews, 1970; Noyes, 1989; Campbell and Hanula, 2007; Fraser et al., 2008; Mazon and Bordera, 2008; Diserud et al., 2013; Schmidt et al., 2019). Within these groups, malaise trapping is especially appropriate for Tenthredinidae, Ichneumonoidea, Scelionidae, Mymaridae, and other hymenopterans with similar life histories, as well as Cicadellidae and Cercopidae (Hemiptera), microlepidopterans (Lepidoptera), and the semi-aquatic orders Plecoptera, Ephemeroptera, and Trichoptera-if traps are placed alongside aquatic habitats. It is important to note that malaise trap efficacy for a taxon can depend on habitat. For example, bees (Apoidea) are sampled well in some habitats, like tallgrass prairie ( Geroff et al., 2014), but pan-trapping is generally more effective for sampling this superfamily ( Campbell and Hanula, 2007). The narrower the taxon of interest, the more necessary it is to customize these recommendations to your own system.

Methodological Considerations

Location

Spatial placement is extremely important for malaise trapping; it is critical to document the trap's exact position and microhabitat by photograph, written description (particularly for habitat), and precise coordinates (see Table 1 for additional metadata needed). Choice of placement will vary based on the particular taxon of interest, but malaise traps should generally be placed along natural flight corridors (e.g., streams or gaps between bushes) to maximize catch. Amount of wind exposure should also be recorded, as wind can limit efficacy. The vertical panel of netting (i.e., the interception area) that insects hit should be oriented to be perpendicular to the expected movement corridor.

Design

We recommend Townes-type malaise traps due to their broad efficacy and already widespread use. Multiple sizes are available, with a 165 × 110 cm interception area being most common. The vertical wall of netting is commonly black to reduce visibility, while the dome of the tent is white to increase insects' propensity to fly upwards (i.e., toward "the sky"), which increases catch. We recommend 95% ethanol (as is used by BIOSCAN) as a killing agent that preserves DNA, but lower concentrations (down to 80%) can be used if evaporation or cost prohibits use of 95%. Alternative approaches to collection include cyanide (hazardous to humans), ethyl acetate (destroys DNA), and live collecting (needs to be checked daily, and specimens are often damaged). Ethanol, however, does remove scales from taxa like lepidopterans. Mesh size and shape are also important to note: holes that are too wide are less effective at sampling small flies, wasps, and other microfauna, but large mesh sizes can better sample groups like stinging hymenopterans ( Darling and Packer, 1988). Some insects, such as many beetles, drop after hitting the mesh screen, rather than trying to escape by flying higher. To take advantage of this, pans with collecting liquid can be placed underneath the mesh wall, creating what is called a flight-intercept trap (traditional malaise traps are a specialized flight-intercept trap that only samples upward-moving insects). If this is done, pan color should be recorded.

Scalability

Commercial malaise traps can be expensive (usually > US $230). Costs can be reduced by constructing home-made malaise traps (e.g., Blackmon, 2010). Mosquito netting can be used for the interception area ( Lamarre et al., 2012), but mesh size and shape should be considered. Setup effort is generally low (up to 1 h the first time). Sampling effort is minimal once a malaise trap is set up (10 min per week once at the site): the sample vial simply needs to be emptied into a storage vial. Post-sampling effort can be high, however, since malaise trapping can yield large numbers of insects (e.g., up to 10,000 specimens per week depending on the site). DNA metabarcoding of the sample is a faster post-sampling identification method for measuring diversity (though abundance is lost), and non-destructive methods to sample from fixative fluids show promise ( Marquina et al., 2019; Nielsen et al., 2019; Zizka et al., 2019). Although metabarcoding lowers the time costs off post-sampling identification, metabarcoding does come with increased costs related to DNA sequencing and additional genetic expertise is needed. See Hausmann et al. (2020) for a recent example of a malaise trap study employing metabarcoding. Finally, because malaise traps are sensitive to microhabitat variation-like most stationary sampling methods-multiple traps at a site are better than a single trap.

Light Trapping

Overview

Light traps are one of the most common and efficient methods for surveying insect that fly at night. At their most basic, light traps simply consist of a light attractant and a viewing surface, often a bedsheet ( Figure 2B). More structured light traps commonly consist of a funnel, vanes (which deflect insects toward the funnel), and a collection container, which together are used in conjunction with the light source to form a structured trap. In either case, light-attracted insects fly toward the light source, hit a surface or vanes surrounding the light, and can then be observed and recorded or sampled and collected. Common styles of vaned light traps include Robinson traps and Heath traps ( Macgregor et al., 2017). Light traps provide an opportunity to gather standardized and comparable data, but many factors influence the abundance and composition of light trapped insects, including trap type, season, time of day, lunar phase, duration of sampling, and light attractant ( Jonason et al., 2014). Consequently, these details are all important to track ( Table 1). Mercury vapor bulbs are the most commonly used attractant and have consistently caught a higher abundance and diversity of insects than other standard bulbs due to the powerful low-wavelength light emitted ( Jonason et al., 2014; White et al., 2016). Other commonly used bulb types include UV, metal halide, and LED ( Ferro and Summerlin, 2019). Although many commercial light traps are available and can be deployed in remote locations, light trapping can be as simple as documenting the moths that are attracted to your porch light. Individuals interested in joining the Discover Life's Mothing project can join a network of people working to photograph and identify moths that come to their porch light ( Pickering, 2015). Exemplar long term light-trapping programs include the Hungarian Light-trap Network ( Szentkirályi, 2002) and the Rothamsted Insect Survey ( Macgregor et al., 2019), both of which have been surveying phototactic insects for over 50 years.

Taxonomic Considerations

Methodological Considerations

Location

Spatial placement is extremely important for light trapping. Although light traps can attract insects from the surrounding environment, insects are rarely attracted at distances >30 meters ( Truxa and Fiedler, 2012). Therefore, the microhabitat of the trap location will influence what organisms are trapped, making it important to describe the trap location in field notes and record the precise coordinates of the trap location ( Table 1). Light pollution can decrease the flight-to-light behavior of moth populations ( Altermatt and Ebert, 2016), so light sensors (low quality light sensors are available as smart phone apps) should be used to note the lumens/m ² of light pollution at the trap location.

Design

The wavelength and brightness of light attractants differs dramatically among different light bulbs and are important to consider when designing light trapping projects. If the goal is to sample the greatest abundance and diversity of insects, then we recommend using mercury vapor bulbs, as these are consistently found to attract the most moths ( Jonason et al., 2014; White et al., 2016). However, mercury vapor traps may not be the optimal tool because of their cost and the logistics required to deploy them (e.g., an outlet or automotive battery is needed). Therefore, low-cost, light-weight, and easy to deploy light traps offer convenient alternatives and facilitate insect trapping at more sites and in more diverse settings ( White et al., 2016). If light traps can be checked early in the morning, insects can be trapped alive by having traps lined with egg cartons to provide areas for the insects to hide ( Jonason et al., 2014). Live specimens can be photographed in the field or later in the lab, after cooling in a refrigerator and then released the following night ( Ford et al., 2020). If observation alone is undesirable, light trap containers can be lined with pest strips (18.6% dichlorvos [2,2-dichlorovinyl dimethyl phosphate]) or filled with ethanol.

Scalability

Commercial light traps can be expensive (between US $75 and US $500). Additionally, commercial light traps often require outlets or car batteries, making carrying them into remote locations challenging. Homemade light traps can be inexpensively constructed and can greatly reduce weight by running efficient LED strips using small 12V batteries ( White et al., 2016). Effort required for field sampling can be high if insects are live trapped and identified in the field or transferred to jars to be photographed. If insects are lethally trapped, sampling effort can be low, with just a few minutes spent setting the light trap each sampling event and a few minutes spent collecting the specimens the following morning. However, post processing costs can be high, as light traps can yield a high diversity and abundance of insects. Technological advances in "smart light traps," where insects attracted to lights are automatically photographed throughout the night, offer great promise to increase the scalability of light trap surveys ( Hogeweg et al., 2019).

Pan Trapping

Overview

Pan traps ( Moericke, 1951) are trays filled with liquid set out to collect insects. Pan traps often rely on color as an attractant and are effective primarily because insects mistake them for food resources. An insect flies to a pan, attempts to land, then becomes trapped in the liquid solution-often soapy water, propylene glycol, or saline ( Figure 2). Pan traps can be made from nearly any object that holds liquid-i.e., a disposable plate filled with water and a few drops of dish soap-and this accessibility has made them more popular than more training-intensive methods that may sample more diversity (e.g., standardized sweep-netting; Cane et al., 2000). Like all sampling methods, there is no doubt that pan traps have considerable sampling bias for certain taxa ( Portman et al., 2020). For detailed accounts of history and methodology, see Droege et al. (2017), LeBuhn et al. (2003), Vrdoljak and Samways (2012), and Southwood and Henderson (2000). An exemplar pan trap monitoring networks is the UK Pollinator Monitoring Scheme, and a data-recording scheme designed for bee monitoring can be found in LeBuhn et al. (2003).

Taxonomic Considerations

Pan trapping is appropriate for monitoring flying insects ( Figure 1C). It is effective at sampling aphids (Hemiptera), thrips (Thysanoptera), bees and parasitic wasps (Hymenoptera), flies (Diptera), some beetles (Coleoptera), and even some grasshoppers (Orthoptera) ( Evans and Bailey, 1993; Westphal et al., 2008; Vrdoljak and Samways, 2012). Trap efficacy for each taxon varies strongly with pan color ( Vrdoljak and Samways, 2012). Yellow is most commonly used, as yellow traps often collect the largest catches and highest total insect diversity, but other common colors include blue, white, red, and green. As with other monitoring methods, habitat and geographic region can affect the trap efficacy for a given group ( Vrdoljak and Samways, 2012; Saunders and Luck, 2013). For those with broad taxonomic interests for their monitoring programs, we recommend what has become a common standard: yellow pan traps in conjunction with white and blue pan traps ( Vrdoljak and Samways, 2012; Sircom et al., 2018), as is done in the UK Pollinator Monitoring Scheme. If needed, traps can be painted using colors defined by the Bee Inventory Plot program ( LeBuhn et al., 2003).

Methodological Considerations

Location

Pan traps are typically placed in open areas where they can be seen by target insects. Traps can be placed together as close as 5 m-the minimum distance at which they do not influence each other ( Droege et al., 2017). Although a large diversity of spatial arrangements exist, we recommend one of two methods. The first method, used by the UK Pollinator Monitoring Scheme, places 1 trap per square km, and is suitable for sampling large geographic regions. The second method, from the Bee Inventory Plot ( LeBuhn et al., 2003), uses 15 traps in a single array, each separated by 5 m and placed in two perpendicular lines forming an "X." This arrangement is suitable for targeted monitoring, with two arrays (30 traps) demonstrated as being adequate for sampling local bee diversity ( Shapiro et al., 2014).

Design

For discussion of pan trap color, see Taxonomic considerations. Pan traps can be placed on the ground (most common), elevated above the ground, or placed flush with the substrate (i.e., essentially modified pitfall traps). Elevated pan traps sometimes yield larger numbers of specimens ( Tuell and Isaacs, 2009), and pan traps flush with the ground can also attract ground-dwelling species ( Ernst et al., 2016). Trap size may not affect catch ( Gonzalez et al., 2020), so small pan traps are desirable to minimize costs; circular pans with a 7 cm diameter are common. In arid areas, the trap solution may evaporate too quickly between visits, so larger pan traps can be used, for example, 2-gallon buckets. The amount of liquid in a trap can affect trap efficacy and should be recorded when setting up and checking traps. We recommend premixing the liquid solution recipe of LeBuhn et al. (2003): 1 part dish soap to 750 parts water (approximately 1 teaspoon soap for a gallon of water).

Because pan color is the main attractant, it is important to maintain trap color through frequent cleaning and eventual trap replacement when color fades. Scented water like rose water can be used to increase catches for some taxa ( Laubertie et al., 2006) but since maximizing the number of individuals caught is not necessarily a goal of standardized monitoring, we recommend against using scents or baits for benchmarking. Small amounts of preservative chemicals can be added to prevent fungal growth when the time between visits is necessarily long, but chemical safety precautions should be taken.

Scalability

Pan traps are low cost but require frequent (often daily) trap visits and maintenance. For an estimated time budget for a 24-pan transect, see Droege et al. (2017). Specimen processing times can be high depending on target taxon abundance. A typical pan in a field could yield only a few specimens over 24 h, but even with low abundances, numbers can rise quickly if using multiple traps over long time periods. Sieving with nylon mesh (e.g., an aquarium net) is a common practice that speeds up specimen processing, but can damage small, fragile taxa such as aphids.

Pitfall Trapping

Overview

Pitfall traps ( Hertz, 1927) are containers placed flush with ground level to capture ground-dwelling (epigeic) insects. In essence, an insect walks to the trap edge, loses balance, and falls in ( Figure 2D). The container is then checked, the catch collected or documented, and reset. Several recent reviews have discussed pitfall trapping ( Skvarla et al., 2014; Brown and Matthews, 2016; Hohbein and Conway, 2018), and standardized traps have been proposed by Brown and Matthews (2016). Like any insect sampling method, pitfall traps produce taxonomically biased samples ( Topping and Sunderland, 1992; Lang, 2000), but are inexpensive and popular for monitoring. For detailed methodological accounts, see Southwood and Henderson (2000), Brown and Matthews (2016), and Hoekman et al. (2017). Existing pitfall trap monitoring networks include the US National Ecological Observatory Network (NEON; Hoekman et al., 2017), and the UK Environmental Change Network ( Brooks et al., 2012).

Taxonomic Considerations

Methodological Considerations

Location

Pitfall traps can be placed nearly anywhere with suitable substrate for digging. There is some controversy over how far apart traps should be placed; some studies have found that traps provide independent samples even when only 1 m apart ( Ward et al., 2001), while others recommend 10 m ( Hohbein and Conway, 2018). NEON, which conducts standardized trapping across North American sites, separates traps by at least 25 m. Digweed et al. (1995) found that population depletion occurs when traps are separated by 10 m or less, but not if separated by distances of <25 m. Until there is more consensus, a 25 m distance between traps should be adequate to ensure independence of samples.

Design

Pitfall traps can be made of glass, plastic, or metal, but disposable plastic cups have become perhaps the most widespread trap container. The container should be placed with its lip flush with the soil surface. As might be expected, the diameter of the trap affects catch ( Abensperg-Traun and Steven, 1995). Collecting fluid should generally be used to avoid damage to specimens from other trapped insects and to prevent escapes, but the type of collecting fluid used can affect the taxa attracted ( Skvarla et al., 2014). Ethylene glycol has been traditionally used but is toxic for wildlife if consumed and can be easily substituted with propylene glycol, which we recommend for most uses. Propylene glycol evaporates more slowly than ethanol and adequately preserves most DNA, at least over the short-term ( Nakamura et al., 2020). Traps with baits will be more readily disturbed by vertebrates ( Vandenberghe, 1992), and should be avoided. Fences, or guidance barriers that direct insects toward the trap, can increase catch ( Boetzl et al., 2018) but require more effort to set up. Pitfall color affects taxonomic composition of the catch ( Buchholz et al., 2013) and should be recorded; we recommend using transparent containers as described in Brown and Matthews (2016). Using funnels increases catch efficiency while simultaneously reducing vertebrate bycatch ( Radawiec and Aleksandrowicz, 2013), although low roofs also reduce vertebrate bycatch ( Hoekman et al., 2017). Roofs also prevent rain from diluting the collecting fluid and appear to not influence the composition or magnitude of insect catch ( Buchholz and Hannig, 2013). Containers should be nested to allow fast and easy removal of samples. Disturbance from trap placement can affect catch, so a latent period of 1-2 weeks before trap monitoring begins should be observed if possible ( Greenslade, 1973).

Calls for pitfall trapping standardization have a longer history than other monitoring methods ( Brown and Matthews, 2016), and we recommend alignment with existing programs. Given the broad extent of NEON, we recommend that new monitoring programs (at least those in North America) adopt the NEON pitfall trapping protocol when practical ( Hoekman et al., 2017). This protocol involves nested clear plastic cups (diameter: 11 cm, depth: 7 cm, volume: 473 mL) and a roof made of hard plastic raised 1.5 cm above the trap entrance. Each trap is filled with 150 mL of an equal ratio of propylene glycol to distilled water. Traps are placed in arrays of 4, arranged in a square with sides 25 m long.

Scalability

Pitfall trapping using disposable plastic cups is relatively cheap and easily scalable. Set-up can be labor intensive depending on the design (and the inclusion of fences), but under the NEON protocol is limited to simply digging an appropriately sized hole and placing the trap and roof. Checking traps is also a low time commitment, especially when using a nested cup design, which allows for easy removal.

Beating Sheets

Overview

A beating or beat sheet is a piece of fabric supported by a frame, which is placed below a substrate of interest (e.g., a tree branch). An insect rests or feeds on the substrate, which is then shaken or hit ("beat"), dislodging the insect so that it falls on the sheet where it can be collected or recorded ( Figure 2E). This active sampling method is often used in conjunction with an aspirator to suck up fast-moving taxa. Currently, the most common design is two pieces of wood or PVC pipe forming an "X," with a piece of white fabric (e.g., bedsheet) stretched behind. Alternative designs include simply placing a sheet on the ground, or even using an umbrella. Beating sheets are cheap, easy to build, and straightforward to use. Insects can be recorded visually or collected for further identification depending on project goals. The Caterpillars Count! citizen science program ( Hurlbert et al., 2019) is an example of a beating sheet monitoring network in North America.

Taxonomic Considerations

Beating sheets are appropriate for sampling tree and shrub dwelling insects, such as caterpillars (Lepidoptera), some true bugs (e.g., aphids and scale insects; Hemiptera), some beetles (Coleoptera), and other plant-feeding insects ( Figure 1). It is not a good method for sampling flying insects; they will often fly away when the branch is hit, or hit the sheet, then quickly escape.

Methodological Considerations

Location

Beating sheets can be used anywhere vegetation is found for beating: typically shrubs and trees, but also groundcover in some cases.

Design

The standard size and shape for a beating sheet is a square with sides of about 90 cm (3 feet), using two pieces of PVC or wood 1.3 m (51 inches) long for crossbars. A cloth can then be stapled or glued to the crossbars. White cloth should be used to maximize visibility of insects that fall on the sheet. The object used for hitting the substrate can vary, but dimensions should be recorded; a stick of about 2.5 cm (1 inch) diameter and 60 cm (2 feet) long works well. The surveyor should strongly hit the branch 10 times, but not so strongly that the plant is damaged. Because many insect species have some degree of host specificity, substrate type (e.g., tree species) strongly predicts insect species diversity and abundance. This will affect sampling decisions depending on your goals; sampling plants of only one species or of multiple species are both reasonable. Either way, we strongly recommend always recording the plant species ( Table 1). If collecting specimens, using an aspirator helps capture fast-moving or flying insects.

Scalability

A simple beating sheet can be constructed in <15 min using materials that cost < US $10. As an active sampling method, using beating sheets can be more time consuming than passive methods, but usually not prohibitively so. Sampling a substrate and collecting the specimens can take <5 min. Visual surveys can be substituted for specimen collection to reduce time spent on post-sampling identification, but at the likely cost of taxonomic resolution. Only one beating sheet is needed per person sampling. Some research suggests that three plants of the same species is the minimum necessary to accurately estimate insect abundance ( Harris et al., 1972).

Acoustic Monitoring

Overview

Acoustic monitors provide a passive, non-destructive method to detect and identify insects ( Ganchev et al., 2007; Mankin et al., 2011). Insects may generate bioacoustic signals as a means of communication ( Penone et al., 2013), or as a by-product of locomotion ( Kawakita and Ichikawa, 2019). These bioacoustic signals may be captured as sounds with microphones or as vibrations with contact sensors ( Figure 2F). Although contact and ultrasonic sensors have been successful in detecting insect pests that live inside agricultural products ( Mankin et al., 2011), we focus here on acoustic recording units and their use for surveying the relative abundance and diversity of insects. Many factors influence the efficacy of acoustic devices in identifying and estimating density of insects, including the frequency range, substrate (air or water), type of sensor, the size and behavior of the insect, and the distance between the insects and the sensors ( Gibb et al., 2019). These factors should be considered and noted when conducting surveys using acoustic monitoring. Large-scale acoustic monitoring has been successfully coordinated by the French National Museum of Natural History to assess the impacts of anthropogenic stressors on Orthoptera communities ( Penone et al., 2013; Jeliazkov et al., 2016).

Taxonomic Considerations

Acoustic monitoring is an appropriate method for monitoring insects that use sounds or vibrations in communications. Some of these noises, such as cicada and cricket songs can be detected over long distances. If the bioacoustic signal produced by insects follows a consistent species-specific pattern, it can be extracted from background noise for identification purposes ( Ganchev et al., 2007). Therefore, passive acoustic monitoring is particularly well-suited for loud terrestrial insects such as Orthoptera or Cicadoidea because they produce species-specific mating calls ( Penone et al., 2013) ( Figure 1), but may also be useful for a variety of other insects including bees ( Galen et al., 2019; Kawakita and Ichikawa, 2019) and aquatic Hemiptera ( Desjonquères et al., 2020; Gottesman et al., 2020).

Methodological Considerations

Location

The distance at which acoustic signals can be detectable above ambient noise varies depending on the sound's amplitude and frequency, landscape heterogeneity such as topography and vegetation, and weather ( Gibb et al., 2019). Additionally, anthropogenic sounds or sounds from other animals can mask target sounds. The precise GPS coordinate of where static sensors are deployed must be recorded and potential sources of sound pollution should be noted ( Table 1). Bioacoustic devices can also be used while traveling along transects, but we recommend keeping bioacoustic devices in fixed locations to collect data that is easier to standardize across sites and replicate across visits.

Design

Commercially available acoustic monitors can be flexibly programed to collect acoustic signals and on-board metadata for long intervals across a variety of sampling regimes ( Hill et al., 2019). The use of inexpensive components (e.g., microelectromechanical systems microphones) may decrease financial barriers to initiating multisensor surveys but can lower data quality by having lower signal-to-noise ratios and inconsistent frequency response ( Gibb et al., 2019). Critical to successful acoustic surveys is the development of efficient pipelines to process sound files and output annotated data. Manually annotating data is time consuming and can be biased by the analyst's knowledge level. Developing automated machine learning pipelines to process individual sound files-which can each include more than 10 min of ambient sound recording-can both increase the efficiency of data processing while also making data processing more reproducible and interoperable.

Scalability

Recent advances in custom built electronics and the lowering costs of small but usable microphones provide novel opportunities to monitor select insect taxa across greater spatial and temporal scales. Passive acoustic monitors can automatically collect data over long periods (e.g., a month), with minimal maintenance needed to replace batteries and digital memory cards ( Hill et al., 2019). Typically, they are programmed to record periodically during a window of interest. Recent developments in customizable acoustic devices have dramatically lowered costs closer to US $50 ( Hill et al., 2018). Developing automatic identification pipelines using machine learning algorithms is critical to scaling acoustic monitoring and discussed further in section looking forward.

Active Visual Surveys

Overview

Visual surveys are commonly used to document the abundance and diversity of insects that can easily be visually identified in the field, often with the aid of close-focus binoculars and nets ( Figure 2G). These surveys typically involve researchers documenting the presence of a species or counting the total number of individuals of each species observed during a standardized survey. The most frequently used methods include (1) transects, (2) point counts, and (3) area counts. Although mark/recapture is another frequently used visual survey technique to document insect population dynamics, we do not consider that a viable benchmarking technique as it takes enormous effort and would not be tractable to do simultaneously for large numbers of insect species.

All three of the commonly used methods have extensive histories of standardized protocols. Transect counts use visual identification while searching along predefined transects with specified search distances. Pollard walks are a commonly used transect method used in butterfly research, in which an observer visualizes a box that extends 5 m ahead and 5 m to the sides as they walk a transect counting butterflies ( Pollard, 1977). Point counts, where an observer stands still and identifies and counts the number of individuals of the target taxa around them during a set period of time, provides an alternative to transects in sites that are difficult to walk in or where habitats are fragile or at-risk ( Henry et al., 2015). Distance sampling techniques, where observers note their distance from the observed insect, can be implemented with both transects and point counts to estimate densities ( Isaac et al., 2011; Henry et al., 2015). By incorporating imperfect detection, distance sampling allows for density and absolute population size to be estimated in closed populations ( Buckland, 2001). Area counts, such as the North American Butterfly Association's count circle ( Taron and Ries, 2015), consist of surveyors counting each species within a delimited study plot during a certain time period. Insects that are challenging to identify quickly or in flight can be netted and transferred into vials and placed in a cooler to chill until the end of the survey period ( Loffland et al., 2017). Photos of chilled individuals can then be taken for later identification before releasing these individuals. Exemplar visual survey programs include multiple butterfly monitoring schemes (BMS) such as the UK BMS, the Dutch BMS ( Schmucki et al., 2016), and the Ohio Lepidopterists BMS ( Wepprich et al., 2019).

Taxonomic Considerations

Visual surveys are only appropriate for large insects that can be easily detected and identified or photographed in the field. Butterflies (Papilionoidea), dragonflies and damselflies (Odonata), and large bees (Apidae) such as bumblebees are effectively sampled using visual surveys ( Figure 1). When species cannot be identified, individuals can be netted and then identified or photographed ( Loffland et al., 2017; Holtmann et al., 2018). However, not all species can be identified in the field; for some species, microscopic examination of the genitalia and abdominal appendages is necessary for identification. Visual surveys generally focus on generating a complete list of species observed (with or without counts), and therefore, visual surveys must focus on a select target group of insects (e.g., butterflies or bumblebees).

Methodological Considerations

Location

The location of the visual survey is important as many insects are habitat specialists. After selecting sites for visual surveys, multiple transects, points, or areas should be randomly selected to ensure sampling across the heterogeneity of a site. It is important to document the coordinates of the survey location and note the habitat type. Visual surveys occurring in difficult terrain or in at-risk ecosystems may consider choosing point counts to limit trampling or allow for more flexible walking routes.

Design

Scalability

General Methodological Considerations

Replication

Four types of replication are especially important when sampling: spatial, inter-annual, intra-annual, and within-sample replication. All types of replication are important and broaden the inferential scale of any monitoring program while also expanding the analytical options and flexibility. We caution, however, that when sampling lethally, large-scale replication (particularly, spatially and temporally intense sampling at a local scale) could theoretically lead to abundance declines, especially perhaps in rare taxa ( Minteer et al., 2014, but see Gezon et al., 2015). Lethal sampling should only be done when scientifically and ethically justified ( Drinkwater et al., 2019).

The required amount of spatial replication for accurate and precise monitoring is still unknown for many insect monitoring methods, and contentious for others. We provide specific discussion in each method section, but in general, the more spatial replication, the more accurate and precise the estimates of measured responses. One way to increase the degree of spatial replication without increasing individual effort is to join an existing monitoring network, thereby increasing the network's degree of spatial replication.

A high degree of intra-annual replication is also important because insect phenology is complex and variable. For example, the week of peak abundance for a species one year could be different the next year, and even the number of generations produced by a species can vary across years. Even if only interested in studying occurrence or abundance, intra-annual variation in phenology of insect activity and generations can lead to strong biases in these responses if samples are only collected once per year (i.e., the "groundhog effect"; Didham et al., 2020). To account for this, we strongly recommend that monitoring efforts be carried out for the entire season of activity for the taxon of interest, allowing easier comparison across years.

Finally, within-sample replication-that is, multiple samples of a response at a single site during a period when the occurrence and abundance of target taxa are assumed to be constant-can be important for the statistical analysis of trends when trying to account for imperfect detection of individuals and species, such as with occupancy modeling ( MacKenzie et al., 2006). Within-sample replication can be achieved either through spatial sub-samples-for example, using "array" designs, as discussed above for pitfall and pan traps-or temporal sub-samples-such as conducting visual or acoustic surveys at the same location multiple days in a row. In general, the ability to collect within-sample replication of monitoring data depends on the design and scalability of a chosen method. Although within-sample replication can substantially increase effort of monitoring, the ability of increased samples to account for sampling noise can be extremely powerful when detection probabilities of target taxa are low. For recent examples of occupancy modeling using some means of sampling replication for insects, see Isaac et al. (2014), Loffland et al. (2017), Outhwaite et al. (2019), Szewczyk and McCain (2019), and Powney et al. (2019).

Curation of Specimens

Many of the monitoring methods discussed require collecting insect specimens for subsequent identification. After identification, it is generally up to the scientist whether specimens should be kept or discarded, although vouchering of representative taxa can be key to the long-term value of datasets, particularly given taxonomic revisions and new technologies. To that end, great care should be taken to ensure specimens are preserved properly, if kept. For an overview of insect specimen storage, see Heraty et al. (2020). It is becoming increasingly common to preserve specimens in 80-95% ethanol to better preserve DNA. Regardless of the preservation method, it is worth developing a plan for deposition and cataloging of specimens prior to beginning a monitoring project. If discussed ahead of time with university and museum insect curators or collection managers, specimens that are properly preserved and curated should be donated. Optimally, archival deposition should include all survey notes, along with specimens, to enhance long-term re-use value.

Curation of Data and Metadata

Just like physical specimens, data are valuable resources for future science. Just as important as data are metadata, that is, the collective information about how the monitoring data were collected. Each method of insect monitoring has its own unique set of critical metadata ( Table 1). Monitoring programs should strive to meet FAIR data principles to ensure their data and metadata are: findable, accessible, interoperable, and reusable ( Wilkinson et al., 2016). Findable and accessible data will require that collected data are digitized and uploaded to websites or online databases that are constructed to hold data about the collecting method. There is much work on this topic that needs to be done, but some resources are developing rapidly.

Interoperable data may be the most challenging aspect of FAIR principles, given that each monitoring program often develops its own reporting standards, even for the same monitoring methods, which ultimately places the burden on downstream users to reintegrate, often with loss of key information. Recent efforts have called for unified, global monitoring standards, such as the Humboldt Core metadata standard ( Guralnick et al., 2018). While it is unlikely that one data standard will fit all insect monitoring, the Humboldt Core provides a typology of different survey and inventorying processes, such as restricted or open searches, along with key definitions of taxonomic, spatial, and temporal scopes, that strongly aid in discovery of monitoring datasets. More specific metadata describing particulars of different monitoring schemes can and should be accommodated ( Table 1). We argue that rather than assume an improbable utopia of full data integration, monitoring programs should work in federations and take seriously the production of detailed metadata, and, as much as possible, develop standardization for metadata that can be as easily linked as possible into existing frameworks. The end result will be FAIR data that can most easily be integrated into flexible modeling frameworks that allow statistical integration of well-described data to better answer broad-scale ecological questions.

As with specimens, it is critical that a plan for data management be considered prior to beginning a monitoring project. Data standards are a key part of that plan since monitoring metadata is crucial for generating insights from monitoring outcomes. However, other factors are also critical, including developing local data storage solutions, deciding on a longer-term repository for data, assuring appropriate credit models for those involved with data collection, and licensing and use agreements of data products. Each of these issues deserves its own longer contribution and we point readers to Michener (2015) and Hardisty et al. (2019) for further reading. Here we make two broad recommendations. First, we strongly suggest development of a coherent data storage and sharing plan that has community buy-in. A best-case approach is development of an internal content management system that provides tools for data access and curation for program participants along with a broad, coherent, and multipronged, data sharing policy that assures long-term access. One part of this sharing policy should focus on best practices for archiving data in community repositories such as Zenodo and Dryad. A second part should focus on publication to aggregators that specialize in biodiversity data mobilization, such as the Global Biodiversity Information Facility (GBIF). The value of publishing to GBIF is enhanced discoverability, since it acts as a single, global access point to biodiversity data and information. However, it can still be challenging to properly publish all survey metadata given GBIF's reliance on standards that were built for incidental records ( Guralnick et al., 2018). Finally, we also encourage monitoring programs to explicitly state data collection and review policies, including how individuals within the programs are credited for the work they do. Such credit models may include attribution for use of data, which can be supported by both licensing mechanisms such as creative commons licenses, and data use policies.

We also encourage development of digital tools to support the capture of field data. Digital tools (e.g., phone apps) can limit transcription errors and are sometimes easier to manage in the field. However, physical data sheets still play an important role in most monitoring programs and are a reliable backup over the very long-term. For these, archival-quality paper and ink should be used to maximize longevity. Though the cost and effort are not trivial, undigitized data can be digitized increasingly easily via scanning and optical character recognition (OCR) capture. Finally, the entoGEM project ( Grames et al., 2019) is soliciting unpublished insect abundance and diversity time series for inclusion in a global systematic map and meta-analysis. EntoGEM is a database, not a repository, but can serve as a temporary mechanism for archiving until a suitable repository is found and is a way to increase the utility of your data.

Looking Forward

Traditional survey methods are limited by being labor and time intensive, but ecological monitoring of animals has recently undergone a dramatic transformation with the development of technologies that expand the spatial, temporal, and taxonomic scales possible to monitor biodiversity ( Pimm et al., 2015). These technological advances could facilitate the collection and availability of vast quantities of data by reducing the effort and expense of insect monitoring. No single method will be able to monitor multiple different insect groups across diverse landscapes. However, a combination of emerging technologies in surveying methods, processing, and data sharing pipelines will allow insect trends to be extracted at currently unprecedented scales.

Camera traps are another emerging surveying tool that are being used to monitor a variety of wildlife ( Burton et al., 2015). Networks of many camera traps allow for data to be collected across greater spatial and temporal scales ( Kissling et al., 2018). Like with acoustic monitoring, deep learning convolutional neural networks are being developed to automatically count and identify wildlife ( Norouzzadeh et al., 2018). The relatively small size of insects compared to wildlife typically captured using camera traps provides unique challenges to monitoring insects with cameras. However, recent studies have shown the potential of camera traps to monitor the overall abundance of flying nocturnal insects ( Ruczyński et al., 2020). Additionally, a portable computer vision light trap has been developed to attract and identify live moths ( Bjerge et al., 2020), and a monitoring network of camera traps that are made with smart image processing has been proposed to monitor light-attracted insects in the Netherlands ( Hogeweg et al., 2019). Continued effort into developing camera traps designed to monitor insects has great potential for passive surveying of non-acoustically detected insects at greater spatiotemporal scales.

Environmental metabarcoding is an emerging tool that provides rapid and cost-effective means for taxonomic identification of many organisms in terrestrial and aquatic environments ( Piper et al., 2019). These approaches can provide detection/non-detection data for insects collected from a variety of methods, especially if specimens are stored in 80-95% ethanol. Metabarcoding insect feces (e.g., frass) offer another non-lethal surveying tool, as caterpillars have been identified to species by amplifying larval DNA from caterpillar feces ( Rytkönen et al., 2019). Unfortunately, most insects have insufficient reference sequences in public archives such as GenBank, making genetically identifying insects challenging. Still, environmental metabarcoding provides a cost-effective and efficient option to identify insects collected in large quantities.

Radar can also create standardized monitoring data for insects at broad spatial and temporal scales ( Didham et al., 2020). Filtering insects from meteorological data can provide previously unused datasets to monitor insects through time and have been used to document the decline of burrowing mayflies ( Hexagenia spp.) across North America ( Stepanian et al., 2020). In most cases, species-level identification cannot be accomplished using radar approaches, but specialized entomological radar shows promise in monitoring insects that may migrate in large abundances at heights difficult to monitor using traditional approaches.

New methods do not need to work in isolation nor are they replacements for traditional monitoring methods. Rather, these new approaches are ways to augment existing ones, and to lower costs for onerous activities that may be partially or wholly automated. We envision passive monitoring tools that can be deployed in conjunction with traditional trap or restricted search methods. For example, acoustic monitors and camera-loaded light traps could be controlled by one device and augment, for example, pitfall trapping, to capture a broader spectrum of insects at a single site. If these sensor approaches also have means to easily share data across a network of sensors and people, it may speed up necessary steps to create the most usable data resources for broad-scale insect monitoring.

Conclusion

We live in an era of rapid change that affects nearly all life on Earth. We can only understand this change and its effects on insects by pooling effort, integrating projects, and working together. Methods standardization is a relatively simple first step, but what challenges come next? For one, we urgently have to coordinate our use of these tools. Networks of networks need to be built for data collection. This means incentivizing participation, coordinating new and existing projects, and organizing efforts on a trans-national scale. Large networks should communicate with each other to increase complementarity, and smaller networks should seek to fit in with what is already being done while maximizing the utility of their own data. But coordination is likely not the greatest challenge.

The largest bottleneck for insect monitoring is getting from trap to accessible data-we need to accelerate the time-consuming stage spent processing and identifying specimens and build tools for the efficient capture of all data and metadata associated with an observation. Improving identification is an area where we have immense potential for advancement over the near term. Bringing more automation to this stage will result in much shorter lag times between data collection and analysis and increase scalability of new and existing projects. Tools for automating identification include metabarcoding, computer vision, and machine learning. Efficient expansion of these identification tools will not only facilitate the broader participation of individuals in insect monitoring (e.g., those without specific skills in taxonomy), but the digital nature of automated and semi-automated identification will speed up data accessibility and metadata capture.

We also need analytical advances for integrating data collected by multiple sampling methods. Assimilating data collected using the same benchmarking method into a composite database is the first step, but ultimately, integrating data collected by multiple different means will vastly improve our ability to understand the broader insect community. But even if we arrive at the point where all the necessary data are being collected on a global scale, the best data in the world are useless if they are not made available-where availability means in a digital format that adheres to all of the FAIR principles. Consequently, we also need the infrastructure to aggregate, store, and share data widely-existing databases such as GBIF are paving the way-while also recognizing the importance of attributing credit (e.g., Chavan and Penev, 2011) to incentivize participation in the process of infrastructure development and data curation.

Every one of these challenges will require collective action to overcome. In the face of rapid Anthropogenic change, there is an intense urgency to this effort. These are no small tasks, and the timeline for completing them is short. Benchmarking has no point when there is nothing left to benchmark. Appropriate foresight and funding would have developed large-scale insect monitoring long ago, but the second best option is to rapidly build capacity now.

Author Contributions

GM and MT conceived the idea for the manuscript and proposed the initial outline. GM and MB led writing of the first draft. All authors contributed critically to drafts and gave final approval for publication.

Funding

This research was supported by the US National Science Foundation (Grants EF 1703048 and 2033263) to MT and RG. Additional support was provided to GM through the UCLA Department of Ecology and Evolutionary Biology and to MB through a University of Florida Biodiversity Institute Fellowship.

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We thank David Wagner for providing useful comments that significantly improved the manuscript, as well as the valued input of two reviewers.

Footnotes

References

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# data[].attributes.attachment_link.title
Standards and Best Practices for Monitoring and Benchmarking Insects

# data[].attributes.attachment_link.description
Benchmark studies of insect populations are increasingly relevant and needed amid accelerating concern about insect trends in the Anthropocene. The growing recognition that insect populations may be in decline has given rise to a renewed call for insect population monitoring by scientists, and a desire from the broader public to participate in insect surveys.

# data[].attributes.body
Primary literature. A book found in UOW Library that has information on amphibian survey methods.

Dodd, CK 2009, Amphibian ecology and conservation : a handbook of techniques , Oxford University Press, Oxford.

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Australian government department of environment, water, heritage and the arts existing guidelines. Useful because sticking with official government guidelines could help prevent breaches of the law.

http://www.environment.gov.au/system/files/resources/ff3eb752-482d-417f-8971-f93a84211518/files/survey-guidelines-frogs.pdf

# data[].attributes.body
https://www.researchgate.net/publication/248881996_Evaluation_of_the_Stag-Watching_Technique_for_Census_of_Possums_and_Gliders_in_Tall_Open_Forest
This paper evaluates the 'Stag-watching' technique in surveying possums and gliders. "Stagwatching involves direct counts of nocturnal animals emerging from tree hollows at dusk"

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eDNA - cutting edge surveying technique
https://www.ari.vic.gov.au/research/field-techniques-and-monitoring/edna-technology-an-innovative-survey-method

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Amphibian survey and handling guidelines 

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Amphibian survey and handling guidelines 

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https://www.environment.nsw.gov.au/-/media/OEH/Corporate-Site/Documents/Animals-and-plants/Threatened-species/amphibians-field-survey-methods-090213.pdf

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https://www.sciencedirect.com/science/article/pii/S0378112705006092
Research paper on the effects of prescribed fires on roosting sites of evening bat (Nycticeius humeralis)

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Primary source- Use of terrestrial invertebrates for biodiversity monitoring in Australian rangelands

# data[].attributes.subject
Primary source- Use of terrestrial invertebrates for biodiversity monitoring in Australian rangelands

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https://www.ccmaknowledgebase.vic.gov.au/resources/Andersen_et_al_2004.pdf

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https://www.froglife.org/wp-content/uploads/2013/06/Reptile-survey-booklet-3mm-bleed.pdf

# data[].attributes.headline
Primary source- Invertebrate Monitoring in the Corangamite Catchment

# data[].attributes.subject
Primary source- Invertebrate Monitoring in the Corangamite Catchment

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https://www.ccmaknowledgebase.vic.gov.au/resources/InvertebrateMonitoring_FINAL.pdf

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Terrestrial Vertebrate Fauna Survey Guidelines for Queensland Version 3 (www.qld.gov.au)
-Could be used as a step-by-step on how to construct a survey

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https://besjournals.onlinelibrary.wiley.com/doi/10.1111/2041-210X.13258

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Specific Primary Source - arboreal marsupial

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Specific Primary Source - arboreal marsupial

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https://ezproxy.uow.edu.au/login?url=https://www-proquest-com.ezproxy.uow.edu.au/scholarly-journals/passive-acoustic-monitoring-detecting-yellow/docview/2532078172/se-2?accountid=15112

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Sugar Gliders

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https://www.epa.wa.gov.au/sites/default/files/Policies_and_Guidance/Tech%20guidance-%20Sampling-SREs-Dec-2016.pdf

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Limitations of various survey methods regarding arboreal reptiles.

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2018 Barking up the right tree  - WildlRes ACC MS

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https://www.animalethics.org.au/policies-and-guidelines/wildlife-research/pitfall-traps

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https://ecos.fws.gov/ServCat/DownloadFile/124721
Paper on conservation biology. Great starting point for a wide range of techniques, although the paper is a bit older. Concise.

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CSIRO Permit requirements to collect invertebrates

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CSIRO Permit requirements to collect invertebrates

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Links for each state's requirements

https://www.csiro.au/en/about/facilities-collections/collections/anic/permit-requirements

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https://uow.primo.exlibrisgroup.com/discovery/fulldisplay?docid=cdi_proquest_journals_221523746&context=PC&vid=61UOW_INST:ALMA&lang=en&search_scope=MyInst_and_CI&adaptor=Primo%20Central&tab=Everything&query=any,contains,ground%20dwelling%20mammal%20survey%20guidelines&offset=0

# data[].attributes.body
http://www.environment.gov.au/system/files/resources/eba674a5-b220-4ef1-9f3a-b9ff3f08a959/files/survey-guidelines-reptiles.pdf

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https://birdata.birdlife.org.au/survey-techniques

Briefly outlines a few common methodologies for surveying birds

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https://media.australian.museum/media/dd/Uploads/Documents/9382/The+Invertebrate+Collection+Manual.d7d0215.pdf

Outlines traditional methods of surveying invertebrates in Australia.

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https://www.livescience.com/28272-bats.html
Provides summary of types of bats, habitats, diet, lifecycle and conservation

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https://www.epa.wa.gov.au/sites/default/files/Policies_and_Guidance/Tech%20guidance-%20Sampling-SREs-Dec-2016.pdf
Outlines surveying guidelines for a number of invertebrate taxa

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https://www.nature.com/articles/s41597-020-0512-9
Aerial surveys of waterbirds in Australia
Published: 10 June 2020!
Very in-depth description of one technique.

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Wombat surveying using satellite imagery

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Wombat surveying using satellite imagery

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https://uow.primo.exlibrisgroup.com/discovery/fulldisplay?docid=cdi_gale_infotracacademiconefile_A538320007&context=PC&vid=61UOW_INST:ALMA&lang=en&search_scope=MyInst_and_CI&adaptor=Primo%20Central&tab=Everything&query=any,contains,wombat%20survey&mode=basic
Comparing historical imagery and field surveys. Trails and wombat warrens are visible in satellite imagery. Specific to wombat sheltering behaviour.

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Primary source: Innefective/outdated survey guidelines -

# data[].attributes.subject
Primary source: Innefective/outdated survey guidelines -

# data[].attributes.body
https://uow.primo.exlibrisgroup.com/discovery/fulldisplay?docid=cdi_crossref_primary_10_1071_WR17012&context=PC&vid=61UOW_INST:ALMA&lang=en&search_scope=MyInst_and_CI&adaptor=Primo%20Central&tab=Everything&query=any,contains,ground%20dwelling%20mammal%20survey%20guidelines&offset=0

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Forestcheck- ground dwelling species survey techniques

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Forestcheck- ground dwelling species survey techniques

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https://www.dpaw.wa.gov.au/about-us/science-and-research/landscape-conservation-research/183-forestcheck?showall=&start=2

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Existing guidelines
http://www.environment.gov.au/system/files/resources/2f420bf1-d9e4-44ec-a69c-07316cb81086/files/survey-guidelines-bats.pdf
- Survey guidelines specifically for threatened bats in Australia

Primary literature
https://www.jstor.org/stable/3784249
- The comparison of acoustical survey techniques for bats in general

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Primary literature - Invertebrate surverys in NZ & Australia

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Primary literature - Invertebrate surverys in NZ & Australia

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https://newzealandecology.org/system/files/articles/NZJEcol28_1_151.pdf

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https://www.animalethics.org.au/policies-and-guidelines/wildlife-research/wildlife-surveys
NSW Dep. of Primary Industries
Useful overview of ethical considerations when surveying animals in the wild.

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Animal Research Review Panel Guideline 10
Revised: January 2020

These guidelines have been written to provide guidance for members of Animal Ethics Committees (AECs) and investigators who carry out wildlife surveys, including those for Species Impact Statements or Environmental Impact Statements required under environmental protection legislation (Biodiversity Conservation Act 2016, National Parks and Wildlife Act 1974, Environmental Planning and Assessment Act 1979 and the Threatened Species Conservation Act 1995).

There are no general texts that describe the ethical capture and handling of all Australian wildlife in the field, although some have been published overseas (see Section 12). There is detailed recent Australian information on the capture, handling and care of native mammals in the National Health and Medical Research Council (NHMRC 2014) A Guide to the care and use of Australian native mammals in research and teaching and this should be used as a primary reference for this group. More general guidance for all animals is available in NHMRC (2013) Australian code for the care and use of animals for scientific purposes, hereafter referred to as the Code. Information on first aid for injured wildlife can be found in Walraven (1990).

These guidelines are not meant to be exhaustive or prescriptive. Their aim is to provide some general principles for reducing the impact on wildlife during surveys, to promote good practice and to raise issues that AECs need to consider when discussing such proposals. The information they contain is taken from the scientific literature and from animal care statements provided by wildlife surveyors.

1 Definitions

Animal Ethics Committee - A committee constituted in accordance with the terms of reference and membership laid down by the Animal Research Act 1985 and the Australian Code for the Care and Use of Animals for Scientific Purposes (NHMRC).

Anabat ™ detector - An electronic device designed to record the echolocation calls of bats (that are usually beyond human hearing) and analyse the characteristic calls of specific species. There are some limitations to this equipment such as inability to distinguish all species. For example it will not detect flying-foxes as they do not use echolocation. Direct observation can be used for these species.

Box trap - A box made from sheet metal with an open door that is released and closes when an animal interferes with the bait in the trap. Sizes vary from quite small (for catching mice) to traps large enough to bandicoots. Because the most commonly used brand is Elliott, these are often referred to as Elliott traps.

Cage trap - Similar to a box trap except that the trap is made from steel mesh. Sizes vary from quite small (for catching mice) to traps large enough to trap dogs. The most commonly used size is 60 x 30 x 30 cm.

Camera trap - A remotely activated camera that is equipped with a motion sensor or an infrared sensor, or uses a light beam as a trigger.

Direct observation - Standing and watching, or walking in a particular direction for certain lengths of time, using binoculars or a spotting scope to detect the range and number of birds or large mammals.

Elliott trap - See box trap.

Fyke net - A net constructed of hoops decreasing in size with webbing between to form a cone shape and one or more funnels inside that prevent trapped fish from swimming out (ie a 'hoop net'), and which also has wings of one or two pieces of netting at the first hoop which are anchored into position with poles. These wings guide the fish into the net. Also known as wing, frame, trap or hoop nets. When used in waterways with platypus, the end of the net must be elevated above water height so that any trapped platypus or turtles do not drown.

Gill net - A net of diamond shaped mesh that is set vertically. The fish are unable to back out because their gill covers get caught in the mesh. When used in waterways with platypus the bottom of the net must be only lighted weighted so the platypus can swim to the surface if caught.

Hair tubes - Small PVC tubes lined with double sided sticky tape with an internal compartment where bait is placed. They may be more efficient and cost effective than the other methods for some rare or trap shy mammals.

Harp trap - An array of thin nylon fishing lines tensioned between two horizontal poles with an escape-proof hessian pocket located below. Bats fly into the lines, fall down undamaged into the pocket and crawl up to roost under a hessian flap where they can roost until released.

Hoop net - See fyke net.

Mist net - Large very fine nylon nets that are strung across potential flyways close to the ground between the vegetation in order to catch birds or bats that fly into them and become entangled. It is very easy for both birds and bats to injure themselves or become distressed whilst being disentangled from these nets so great care is needed.

Playback calls - Pre-recordings of the calls of nocturnal birds (such as owls), frogs and arboreal mammals (such as the koala) which are then played back at night in order to elicit a response from any member of the target species present which may be a reply (or call back) or an approach. They are usually broadcast at various locations over a specified duration (e.g. 10 minutes initial listening, 15 minutes playing of the recording and 10 minutes listening for a response).

Pitfall trap - A glass, metal or plastic container sunk into the ground so that the mouth is level with the soil surface. Ground dwelling animals fall into the trap and are unable to escape.

Proposal - A written outline of a project put forward for consideration by an AEC.

Spider tubes - Small PVC tubes installed into the ground, covered by a metal or canvas roof. Tubes are checked for sheltering individuals which can be captured by hand for identification.

Trip line - A single nylon line stretched 1.5-3cm above the surface of a body of water where bats are likely to fly, causing bats in flight to fall into the water and swim out where they are captured. These have much greater potential for damage to the animal than harp traps.

Voucher specimen - Any specimen that serves as a basis of study and is retained as a reference. A "type" specimen is a particular voucher specimen that serves as a basis for taxonomic description of that subspecies.

Wildlife - Free-living vertebrates of native, non-indigenous and feral species including captive bred animals and those captured from free-living populations.

2 The Regulation of Animal Care during Wildlife Surveys in NSW

In NSW, the welfare of animals used for animal research is protected by the Animal Research Act 1985. Wildlife surveys are considered as animal research and are therefore subject to the provisions of this legislation. The legislation also requires that the conduct of animal research must be consistent with the provisions of the Code.

Wildlife surveyors and AEC members should be conversant with the animal research legislation and the Code, and particularly Clauses 3.3.33 to 3.3.47 of the Code relating to wildlife and field techniques.

In addition to the legislation and the Code, the Animal Research Review Panel (the Panel) has a number of policies and guidelines relevant to wildlife surveys that should be consulted (see Section 13.0). Investigators should be familiar with the contents of all these documents. Along with general information on the legislation, AECs, the Panel and the Code, the policies and guidelines are available on the Animal Ethics Infolink site.

The intent of the legislation is to protect the welfare of animals and is implemented through the requirement that all animal research must be approved by an AEC before the work commences. The AEC considers the ethical aspects of the study such as the justification for the survey, the methods used and the impact of all procedures on the animals. To assist the AEC in its deliberations, application is made by the investigator on a proposal form that aims to ensure that AEC members have the information they need for their discussions, written in plain English.

Intending wildlife surveyors associated with an Accredited Animal Research Establishment should seek approval from the establishment's AEC. Those not associated with an Accredited Establishment may apply to the Secretary's AEC through the Department of Primary Industries. If the AEC approves the application, the investigator will be issued with an Animal Research Authority for the project.

A Biodiversity Conservation Licence (also referred to as a Scientific Class Licence) issued under Part 2, Division 3, of the Biodiversity Conservation Act 2016 to conduct research is also required from the Office of Environment and Heritage to conduct wildlife surveys in NSW. This licence requires a current Animal Research Authority.

3 General ethical considerations and wildlife surveys

the replacement of animals with other methods;

the reduction of the number of animals used; and

the refinement of the techniques used to reduce the impact on animals.

The Code advances three fundamental concepts for improving the welfare of animals used for scientific research. These are known as the Three Rs and include:

1. Survey planning
Replacement of animals may be possible by assessing the existing recent knowledge of the fauna in the proposed study area and determine if a new survey is both necessary and justified. Sources of such information include:

published records in scientific journals and the newsletters of scientific and natural history societies. If a survey has been done many years ago in the same area a current survey is probably warranted to allow an assessment of possible changes in species abundance;

biodiversity survey reports by the Office of Environment and Heritage, including NSW National Parks and Wildlife Service and Forestry Corporation of NSW;

the Atlas of NSW Wildlife - a whole-of-government system for flora and fauna sightings information;

Birdlife Australia The New Atlas of Australian Birds (Barrett et al. 2004);

Atlas of Living Australia;

previous Species Impact Statements and Environmental Impact Statements available from the Department of Planning and Environment; and

local knowledge (local councils, land management groups, land holders).

Historical records may not be useful in determining current species presence or abundance as these may have changed over time, often due to threatening processes. In cases such as these surveys may have particular value as they may show trends in species distribution and abundance over time.

Reduction in the usage of animals may be possible if the background research suggests recent surveys have been undertaken in an area of interest and therefore a reduction in survey effort may be appropriate. Wildlife surveyors are encouraged to publish survey information whenever possible, and to lodge results where they can be accessed in the future. Examples of such repositories include the Australian Museum and the NSW Wildlife Atlas and the Atlas of Living Australia.

Refinement in survey techniques comes from substituting non-invasive techniques such as camera traps where they can provide the same information as conventional trapping techniques.

2. Survey design and methodology
It is not the purpose of these guidelines to provide detailed technical advice about the design and methodology of wildlife surveys. Rather, the intent is to show how the welfare of animals can be protected during surveys by employing appropriate design and methodology. This section is a general discussion of these issues. In later sections more specific suggestions are made.

The following points should be considered when designing a wildlife survey:

The methods used should be appropriate to the objectives of the project;

The methods used should be based on sound scientific principles so that the results are valid;

The methods used should minimise the impact on the animals; and

Sample sizes should be kept to the minimum required statistically and that number should be able to be justified.

The following general points should be considered when determining the methods to be used in a wildlife survey.

3. Voucher specimens
The collection of voucher specimens is an important part of scientific research. However, it is a practice of concern to some sections of the community. Wildlife surveyors intending to collect voucher specimens should consult the Panel's guidelines for the Collection of voucher specimens. With the recent advances in genetic screening, consider taking only a tissue sample that can be used for genetic analysis. Consult on an appropriate method with a genetic repository such as the Australian Museum or for mammals refer to NHMRC (2014).

Some important points are listed below:

4. Emergency procedures
All applications to an AEC for wildlife research require a detailed description of emergency procedures in the event of injuries to animals, inclement weather, floods, bushfires and the illness or injury of the surveyors. The purpose of these is to ensure that threats to the welfare of animals resulting from emergencies are identified and mitigated.

Investigators should be conversant with the Panel's Emergency Procedures policy. Issues particularly relevant to wildlife surveys include the following.

Arrangements must be made to clear and close all traps in the event of inclement weather, floods and bushfires.

Arrangements must be made to clear and close traps in the event that illness or injury removes the investigator from the field.

Investigators should have the appropriate skills and equipment to euthanase seriously injured animals in the field should this be necessary. Euthanasia must be by an approved method (see section 3.5 below).

Arrangements must be made to appropriately transport seriously injured animals to the nearest veterinarian for treatment, noting that injured animals should be taken to veterinarians initially rather than to wildlife carers.

Any unexpected problems should be reported to the AEC as soon as possible, including mortalities and injuries to animals. Future surveys may need to be modified in the light of these problems.

5. Euthanasia
Emergency euthanasia or killing of specimens for vouchers may need to be carried out by surveyors in the field. The following points about euthanasia need to be considered.

Methods must be humane and produce a painless death as rapidly as possible.

Methods that are acceptable are described in UFAW/WSPA (1989), UFAW (1996, 1997), Reilly (2001) and AVMA (2013). For native mammals the methods referred to in NHMRC (2014) can be followed.

Methods that are not acceptable include car exhaust fumes, cervical dislocation in animals larger than 150g, drowning and freezing. Note that cooling reptiles and amphibians to make them easier to handle is acceptable but, even after cooling, freezing is not an acceptable method of euthanasia.

Surveyors must be trained and competent in the use of the acceptable methods of euthanasia.

Investigators should be conversant with the Panel's policy, The use of restricted drugs and the conduct of restricted acts of veterinary science in animal research, which details the legal requirements for using sodium pentobarbitone.

6. Diseases and parasites

The Three Rs are as relevant to wildlife surveys as to laboratory studies of animals, and in this section examples of their implementation through good survey planning and design and appropriate methodology are discussed. Specific wildlife survey methods are discussed in later sections.

Surveyors should be aware of their potential exposure to zoonotic diseases (those that affect both animals and humans and may be passed between them) that are present in Australian native animals. Some of these diseases can cause serious illness or death. Several examples of zoonotic diseases include:

Australian Bat Lyssavirus - a virus carried by many species of bats that usually causes death unless vaccinated against.

Psittacosis - caused by the bacterium Chlamydia psittaci that is typically carried by birds, especially pigeons and parrots.

Q fever - Caused by exposure to the bacterium Coxiella burnetii that is carried by livestock and wild life.

Salmonellosis - Caused by exposure to the bacterium Salmonella that is carried by reptiles and various other wildlife.

Toxoplasmosis - Caused by the protozoan Toxoplasma gondii that is carried by cats.

Viral and bacterial infections:

mites and ticks - These are known to be carries or various diseases including Australian Tick Typhus, Lyme disease, Babesiosis, tick paralysis and Red Meat Allergy. These infect people when they bite.

External parasites:

cestodes (tapeworms).

nematodes (roundworms).

trematodes (flukes).

Internal parasites:

high levels of personal hygiene;

not eating, drinking or smoking whilst handling animals;

washing field clothes and equipment that has come into contact with animals' blood or body fluids and cleaning all survey equipment between surveys;

basic first aid for treatment of cuts, bites and scratches;

observance of protocols to avoid transmission of frog chytrid fungus as outlined (Department of Environment and Climate Change 2008);

obtaining vaccinations against Australian Bat Lyssavirus before handling microbats and/or flying foxes.

minimise exposure to ticks.

Surveyors should therefore take basic precautions to prevent animal-animal, animal-human and human-animal transfer of disease. Such precautions include the following:

Should anyone who handled animals become ill within two months of a survey, the attending medical practitioner should be informed of the potential exposure to zoonoses. Further information on zoonoses can be obtained from NSW Health.

4 Surveys of terrestrial and arboreal animals.

The Environment, Energy and Science Group has published a number of guidelines on biodiversity surveys and how to sample native fauna. There are also specific guidelines on how to survey for NSW threatened species.

1. Methods not involving animal capture
Some mammal species leave signs (scats and tracks) sufficiently distinctive to provide positive identification. The sign of many Australian mammal species is described in Triggs (1996). Signs that indicate the presence of species or groups of species should be used in surveys wherever possible.

2. Animal signs

3. Hair tubes
The use of hair tubes is described by Scotts and Craig (1988), Lindenmayer et al. (1999) and Mills et al. (2002). Points to consider are:

Ensure that the floor of the tube is free of adhesive tape to prevent small lizards and frogs becoming stuck.

If an animal does become stuck to the tape, do not try to pull the tape off, as this may seriously damage the skin. Either carefully trim the tape on the animal to as small a size as possible (the remaining tape will be shed during normal skin replacement) or gently ease vegetable oil under the tape and slide it off.

Slope hair tubes with the entrance pointing slightly downwards to ensure drainage.

Commercially designed hair funnel traps are available with a specially prepared sticky wafer that minimise accidental captures of animals.

4. Spotlight counts
When spotlighting animals:

avoid prolonged exposure to the light (i.e. more than two minutes);

use a light with a narrow beam; and

when practical, use a red filter or, preferably, a dimmer switch to reduce light intensity for prolonged observations once the animal has been spotted.

Survey with a low wattage light (30 to 50 w) and briefly use a narrow beam of brighter light for identification (SEWPAC 2011a).

5. Camera traps

The Commonwealth Department of the Environment has also prepared survey guidelines for nationally listed mammals, bats, birds, reptiles, frogs and identify techniques are applicable to the species level. See SEWPAC (2010a, 2010b, 2011a, 2011b).

Traps may be infra-red or white flash.

Infra-red cameras are less intrusive but do not distinguish some species sufficiently, as they do not distinguish colour.

Use of a lure, either scent or mechanical, focuses the detection zone and holds the animal long enough to get an image.

Often used in conjunction with other survey methods.

1. Methods involving animal capture

2. Trapping - general

Use the trapping method with the least impact.

Whenever possible, avoid trapping at times of the year when animals may be susceptible to greater stress, such as during breeding seasons or extreme heat or cold. If animals are breeding, minimise their time in traps by checking more frequently and releasing pregnant or lactating females as a matter of priority.

Select the type of trap that is appropriate to the species being targeted.

Ensure all traps are in good working order, cleaned and checked immediately prior to use.

Limit the number of traps set per field worker to that which can be cleared in two hours.

At any one site, unless otherwise justified, limit trapping periods to no more than four consecutive nights with a minimum of three nights between trapping periods to avoid continually trapping the same individuals.

Use a bait appropriate to diet of the target species. The bait should not only lure the animal into the trap, but should also replace the food and moisture it would have consumed had it not been trapped. This is particularly important for small mammals that have high metabolic rates.

Locate each trap to reduce exposure of trapped animals to the sun, wind, rain etc (for example, place traps under shrubs or beside logs).

Avoid placing traps in areas of high ant activity.

Do not trap during periods of badly inclement weather.

Ensure all traps are located and checked each time a trapline is checked and that all traps are removed from the field or closed at the end of the trapping period. If individual traps are numbered and set in order, it makes it easier to ensure that all traps are checked.

For nocturnal species, begin clearing traps at first light and where practical leave the traps closed until late afternoon. During periods of extremely cold weather, cease trapping completely or clear and close traps by 0200 hrs each day.

For diurnal species, have an inspection schedule that minimises the impact on any trapped animals and locate the traps so as to minimise the possibilities of heat or cold stress.

Release animals as soon as possible and where they were caught.

Cease trapping immediately if there has been an unusually high mortality of animals.

1. Box traps (also known as Elliott traps)
In addition to the general points in 4.2.1 above, the following need to be considered.

Provide bedding in the traps. Dry leaf litter and Dupont Hollofill ™ are suitable materials, although the latter sometimes wraps around the animals' feet. Cotton wool should not be used because it absorbs moisture, increasing the risk of hypothermia.

In areas with wetter climates, place traps in a plastic bag, taking care to ensure adequate drainage (slope traps at 10o to the horizontal to allow drainage during rain).

During periods of high temperatures in areas where traps cannot be sheltered from the sun, close traps during the day.

Traps set in trees should be on the opposite side of the tree to the morning sun.

2. Cage traps
In addition to the general points in 4.2.1 above, the following need to be considered.

Set traps in sheltered positions.

Provide shelter for trapped animals by covering the trap with opaque plastic or canvas (cooler areas) or with hessian or shade cloth (hotter areas).

If traps cannot be sheltered from the sun, they should be closed during the day if temperatures are high.

3. Dry pitfall traps
The Panel has developed guidelines for the use of pitfall traps, which should be observed. In addition, consider the following points:

4. Radio-tracking

The use of camera traps has become widespread in fauna survey and applies not only to mammals but also birds and reptiles (SEWPAC 2011a, Meek et al. 2012). Points to consider are:

In general, the following points apply to the use of traps.

1. Methods not involving animal capture
Ultrasound detectors (for example, the AnaBat ™, Song Meter ™) can be used to detect bats without any impact and should be used whenever possible.

2. Methods involving animal capture

3. General

Radio transmitters are rarely necessary for general wildlife surveys so their use is not covered here in depth. See the Panel guideline Radio tracking and GPS tracking.

Whenever possible avoid trapping during the breeding season.

Bats should be released at the point of capture as soon as possible. However, they should not be released in daylight. Those that cannot be released before dawn should be held until the following dusk.

When necessary, bats should be held separately in suspended cloth bags in a dark, quiet and warm place.

Bats may go into torpor in the trap or while held in bags and will need to be re-warmed before release.

Care should be taken when handling both flying foxes and microbats, due to the zoonotic disease Australian bat lyssavirus (see Section 3.6)

1. Harp traps
A description of the use of harp traps can be found in Tidemann and Woodside (1978). Points additional to those in 5.2.1 that need consideration are:

set traps in a sheltered spot in potential flyways;

clear within two hours of dusk and again after dawn but before the sun begins to warm the hessian; and

harp traps must not be used where large numbers of bats could be caught (for example at entrances to roost sites) to avoid the overheating of bats in the collection bag.

2. Mist nets
Points additional to those in 5.2.1 that need consideration are listed below.

3. Trip lines

5 Surveys of bats

Risks of injury to bats, so use other methods whenever possible;

Monitor continually whenever the line is deployed;

Be prepared to enter the water to rescue bats if necessary; and

Have at least one low-powered torch to collect bats since they will swim away from bright lights.

A description of bat survey methods can be found in SEWPAC (2010a). Surveys for bats should be carried out by an experienced bat investigator as (apart from the fruit bats) little is known of their biology or taxonomy and species can be difficult to identify.

The following general points need to be considered when trapping bats:

Before any trapping is undertaken, the likelihood of platypus being present should be ascertained from other sources including recent published reports and observational surveys undertaken by the surveyor.

In deep water habitats, lightly weighted gill nets should be used in case platypus are present to allow them to swim to the surface.

Trapping must not be carried out during the months of October-March, when females are laying eggs and raising young, unless justified to the AEC.

Gill nets should be observed continuously to detect when a platypus is caught, e.g. by the splashing.

Gill nets should be lifted every hour to check for snags and entrapped large fish.

Nets should have a stretched mesh size of at least 150 mm, or be fitted with an entrance grill with a mesh size of less than 35 mm, or be set so that there is an air space available along the length of the net.

Due to the potential to cause deaths of platypus and turtles check with NSW Fisheries before deployment. Note that it is illegal to use opera-house style traps in some NSW waterways.

Captured platypus should be transferred to a clean bag or secure box and held in a quiet place until all nets are closed for the night, to minimise the likelihood of hypothermia, attack by predators or by other platypus (particularly between males in the breeding season).

Note that trip lines are ineffective for bats that can take off from water (e.g. fishing bats).

Points additional to those in 5.2.1 that need consideration are:

1. Methods not involving animal capture

Avoid close range inspection during breeding and feeding.

Carry out searches for nests, mounds, display areas, characteristic scrapes and scratchings, visual and auditory searches such as breeding calls.

2. Direct Observation

3. Playback calls

Avoid prolonged exposure by limiting calling sessions to two 15 minute periods per night.

Use of play back calls during the species' breeding season should be done with care so as not to disrupt the breeding of the resident animals.

4. Acoustic Monitoring
The advent of cheap digital acoustic recorders that allow the continuous recording of ambient noise has allowed acoustic monitoring to become a viable method of survey. Subsequent sound analysis, using specialised software or expertise, identifies the calls of particular species. This could be an efficient method of detecting species that only occasionally frequent an area or are highly secretive and difficult to detect.

5. Spotlighting owls

6 Platypus

Platypuses are one of the most difficult species to handle and maintain in captivity, because they are highly susceptible to stress unless managed carefully and with a high degree of knowledge and expertise (NHMRC 2014). Wildlife surveys would rarely need to capture platypus to establish their presence. Direct observation at dawn and dusk of suitable riverine habitat is sufficient to establish their presence but not their absence.

7 Surveys of birds

Because of the high risk of injury and death to birds, mist nets should only be used where other methods have already been rejected as unsuitable.

Mist nets must only be used by trained and competent personnel. An appropriate authority for the use of mist nets by the Australian Bird and Bat Banding Scheme (ABBBS) is preferred.

Mist nets should be attended at least every 30 minutes and captured birds removed immediately.

Nets must be closed when not attended.

Numerous methods have been developed for the detection of birds in wildlife survey and the structuring of sample design (SEWPAC 2010b). Choosing an appropriate design and method will depend entirely on the questions being asked.

1. General
A summary of survey methods for reptiles can be found in Blomberg and Shine (1996).

Choosing the correct season is critical for effective surveys of frogs (and to a lesser extent with reptiles). Most frog species are active only during the warmer months of the year (Spring-Summer-Autumn), although there are some that are active only during the cooler months (Autumn and Winter). Outside of their active season many frogs aestivate or go into torpor, usually in burrows, hollows in trees, crevices in timber or rocks or under loose soil. When in torpor, they are undetectable. To a lesser extent this may also occur during the active season when weather conditions are unsuitable (e.g. dry).

2. Surveys not involving animal capture

3. Spotlighting frogs with or without using playback

Avoid excessive foot traffic around the water body.

Keep exposure to a minimum to prevent overheating.

Use a lower intensity light held at a distance for further observations.

1. Camera Traps
Camera traps can be deployed for reptiles when mounted vertically over a gap in a drift fence. This works particularly well for larger lizards and snakes that are not readily captured with conventional means. See Welbourne (2013, 2014) for this technique.

2. Surveys involving animal capture

3. General

1. Freshwater turtles

Set traps with an air space to prevent drowning of turtles or by-catch such as platypus, water rats or water birds. The air space can be maintained by use of a float (e.g. an empty drink container) or by tying the trap to an overhanging tree or log. Opera-house style traps can be tied to a stake on the bank. Note that is illegal to use opera-house style traps in some NSW waterways; check with NSW Fisheries before deployment.

Traps should be checked at least at dawn and dusk. They should be checked more frequently if turtle numbers are high and during summer.

Transport animals separately to avoid the risk of shell damage and hence infection. Keep cool during transport to avoid heat stress.

2. Marine turtles

Marine turtles are very susceptible to heat stress, especially during transport. They can be cooled by the use of wet hessian bags.

Confining the animals in small spaces increases the risk of abrasions, and hence infections. Marine turtles are best restrained by placing them on their backs in a cool place.

During transport, insulate from heat and also from vibration. They are best transported within a vehicle rather than in the tray of a utility.

Consider that hand searches carried out by experienced personnel under suitable conditions will locate nearly all species of reptiles and frogs in an area within a short period of time that may mean that fewer traps or no traps at all need be set.

Frogs should be handled as little as possible because handling removes skin secretions and predisposes the frog to fungal infections (White 1990), while continuous holding in the hand can result in overheating.

Hygiene precautions as detailed in Department of Environment and Climate Change (NSW) (2008) must be observed when handling frogs and tadpoles, including the use of gloves.

Gloved hands should be made wet in the local water or in wet grass/vegetation so that loss of skin secretions is minimised when frogs are first picked up.

Frogs should be moistened with rainwater or water from the stream being surveyed after holding. They may be held separately temporarily (up to 24 hours) in a new moist plastic bag containing some vegetation (although, in the dark, vegetation will absorb oxygen).

Reptiles should be held separately in appropriately sized secure bags or boxes with some vegetation, or a moist paper towel, as appropriate, in a cool place.

Tadpoles are often easier to find than adult frogs and provide important information about habitats used and other measures of environmental quality. However, care needs to be taken when handling tadpoles, as handling can result in a high level of injury and death of the tadpoles. See Anstis (2002) for identification keys.

Consider that fish are usually in their best condition in spring and early summer and will be able to cope with the shock of capture and recover more quickly than in the winter or in mid-summer after spawning.

Use nets with soft mesh (for example, cotton or nylon) to reduce damage to the fish.

Use appropriately sized and weighted traps to reduce the risk of non-target animals being caught.

Fyke nets should have an air space by being set partially out of the water to prevent drowning of trapped mammals (such as platypus and water rats) or waterfowl. Otherwise they should have a means of escape.

If possible, avoid using gill nets because fish caught in these often die (or are so damaged during removal that they are unlikely to survive) and because they have the potential to trap many non target species.

Check and empty traps regularly.

Handle the fish as little as possible.

Minimise the removal of the fish's protective mucous covering and reduce temperature shock by wetting hands first in the water that the fish was caught.

If electrofishing is being used for sampling, operators should have appropriate training and follow guidelines set out in NSW Fisheries (1997).

1. Hand searching for amphibians and reptiles

Take care when moving and repositioning rocks and logs to prevent animal injuries and avoid causing habitat disturbance that may affect the subsequent abundance of the species.

Wash hands without soap (for instance in the water of the water body being surveyed or with rainwater) to reduce contamination from chemicals.

Noose type devices to catch large reptiles should be used with care and devices used to pin snakes need to be padded to avoid causing injury.

2. Pitfall traps
Note that the Panel's guideline Use of pitfall traps should be followed.

See also section 4.2.4 Dry pitfall traps.

3. Spider tubes

Examples of techniques to census owls can be found in Kavanagh and Peake (1993).

See also section 4.1.3 Spotlight counts

Guidelines for using mist nets can be found in the Australian Bird Banders Manual (Lowe 1989).

8 Surveys of reptiles and frogs

Some species of reptile including Grassland Earless Dragon inhabit burrowing spider tubes. Inspection of these visually or with an endoscope can be used to detect the presence of these species.

9 Surveys of turtles

10 Surveys of fish
Department of Environment and Climate Change (NSW) (2008) Hygiene protocol for the control of disease in frogs. Information Circular No. 6. DECC (NSW). Sydney South
General information on survey design and best use of methods is available in Merrick (1990), SEWPAC (2011b) and Barker et al. (2009).
Fair, J., Paul, E. & Jones, J. (Eds) (2010) Guidelines to the use of wild birds in research. Ornithological Council, Washington DC.
11 References

American Society of Ichthyologists and Herpetologists (2004) Guidelines for the use of live amphibians and reptiles in field and laboratory research.Second Edition. Herpetologists' League, Society for the Study of Amphibians and Reptiles. 42pp.

American Veterinary Medical Association (2013) AVMA Guidelines for the euthanasia of animals: 2013 edition. American Veterinary Medical Association, Schaumburg, Illinois.

Anstis, M. (2002) Tadpoles of South-east Australia. New Holland Publishers, Sydney.
Meek, P. & Fleming, P. (eds.) (2014) Camera Trapping: Wildlife Management and Research. CSIRO Publishing, Melbourne.
Bali, R. & Delaney, R. (1996) Assessment of Koala Radiocollaring Studies in South eastern Australia. NSW National Parks and Wildlife Service, Sydney.
National Health and Medical Research Council (2013) Australian code for the care and use of animals for scientific purposes, 8th edition. Canberra: National Health and Medical Research Council. National Health and Medical Research Council (2014) A Guide to the care and use of Australian native mammals in research and teaching. National Health and Medical Research Council, Canberra. NSW Fisheries (1997) Australian Code of Electrofishing Practice. NSW Fisheries Management Publication No. 1
Barker, D., Allan, G.L., Rowland, S.J., Kennedy, J.D. & Pickles, J.M. (2009) A Guide to Acceptable Procedures and Practices for Aquaculture and Fisheries Research. Primary Industries (Fisheries) Animal Care and Ethics Committee, Nelson Bay NSW.

Barrett, G., Barry, S., Cunningham, R., Poulter, R. & Silcocks, A. (2004) The New Atlas of Australian Birds. CSIRO Publishing, Melbourne.

Blomberg, S. & Shine, R. (2006) Reptiles, pp. 297-307, in J. Sutherland (ed.) Ecological Census Techniques: A Handbook. Second Edition. Cambridge University Press, Cambridge.

DeTolla, L.J., Srinivas, S., Whitaker, B.R., Andrews, C., Hecker, B., Kane, AS. & Reimschuessel, R. (1995) Guidelines for the Care and Use of Fish in Research. ILARJournal 37: 159-173.

Helman, P. & Churchill, S. (1986) Bat capture techniques and their use in surveys. Macroderma 2: 32-53.

Kavanagh, R. & Peake, P. (1993) Survey procedures for nocturnal forest birds: an evaluation of the variability in census results due to temporal factors, weather and technique. Pp. 86-100, in P. Olsen (ed.) Australian Raptor Studies. Australian Raptor Association, RAOU, Melbourne.
Sikes, R.S. & Gannon, W.L. and the Animal Care and Use Committee of the American Society of Mammalogists. (2011) Guidelines of the American Society of Mammalogists for the use of wild mammals in research. Journal of Mammalogy 92: 235-253. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2010a) Survey guidelines for Australia's threatened bats. Commonwealth of Australia, Canberra. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2010b) Survey guidelines for Australia's threatened birds. Commonwealth of Australia, Canberra. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2011a) Survey guidelines for Australia's threatened mammals. Commonwealth of Australia, Canberra. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2011b) Survey guidelines for Australia's threatened fish. Commonwealth of Australia, Canberra.
Lindenmayer, D.B., Incoll, R.D., Cunningham, R.B., Pope, M.L., Donnelly, C.F., MacGregor, C.I., Tribolet, C. & Triggs, B.E. (1999) Comparison of hairtube types for the detection of mammals. Wildlife Research 26: 745-753.

Lowe, K.W. (1989) The Australian Bird and Bat Banders Manual. Australian National Parks and Wildlife Service, Canberra.

Mills, D.J., Harris, B., Claridge, A.W. & Barry, S.C. (2002) Efficacy of hair-sampling techniques for the detection of medium-sized terrestrial mammals. I. A comparison between hair-funnels, hair tubes and indirect signs. Wildlife Research 29: 379-387

Parnaby, H. (1992a) An interim guide to the identification on insectivorous bats of southeastern Australia. Technical Reports of the Australian Museum No. 8. Australian Museum, Sydney.

Parnaby, H. (1992b) Summary of the ultrasonic survey of the microbats of north east New

South Wales, 1991/1992. Report to the NSW National Parks and Wildlife Service, Sydney.

Petit, S. & Waudby, H.P. (2012) Standard operating procedures for aluminium box, wire cage, and pitfall trapping, handling, and temporary housing of small wild rodents and marsupials. Wildlife Research 60: 392-401.

Reilly, J. (ed) (2001) Euthanasia of animals used for scientific purposes. ANZCCART, Glen Osmond, South Australia.

Scotts, D.J. & Craig, S.A. (1988) Improved hair sampling tube for detection of rare mammals. Australian Wildlife Research 15: 469-472.

Tidemann, C.R. & Woodside, D.P. (1978) A collapsible bat trap and a comparison of results obtained with the trap and mist nets. Australian Wildlife Research 5: 355-362.

Triggs, B. (1996) Tracks Scats and Other Traces: A Field Guide to Australian Mammals. Oxford University Press, Melbourne.

Universities Federation for Animal Welfare (1996) Recommendations for euthanasia of experimental animals: Part 1. Laboratory Animals 30: 293-316.

Universities Federation for Animal Welfare (1997) Recommendations for euthanasia of experimental animals: Part 2. Laboratory Animals 31: 1-32.

Universities Federation for Animal Welfare /World Society for the Protection of Animals (1989) Euthanasia of Amphibians and Reptiles: Report of a Joint UFAW/WSPA Working Party. UFAW, South Mimms, Potters Bar, Herts, UK. And WSPA, London, UK.

Van Dyck, S. & Strahan, R. (2008) The Mammals of Australia. New Holland Publishers, Sydney.

Walraven, E. (1990) Taronga Zoo's Guide to the Care of Urban Wildlife. Allen and Unwin, Sydney.

Welbourne, D. (2013) A method for surveying diurnal terrestrial reptiles with passive infrared automatically triggered cameras. Herpetological Review 44: 247-250.

Welbourne, D.J. (2014) Using camera traps to survey diurnal terrestrial reptiles: a proof of concept. pp. 225-232, P. Meek, P. Fleming, G. Ballard, P. Banks, A. Claridge, J. Sanderson & D. Swann D (eds.) Camera Trapping Wildlife Management and Research. CSIRO Publishing, Melbourne.

Acknowledgments

Stephen Jackson (NSW DPI) for the development of the current edition and Mike Fleming (DPIE) for providing comments on the draft document. The previous edition of these guidelines was developed with the assistance of M. Murray, P. Burcher, A. Adair, K. Kendall, Leong Lim, ERM Mitchell McCotter, Bob Harden, Gerard Körtner, Fritz Geiser the Australian Platypus Conservancy, the Victorian Department of Natural Resources and Environment, members of the Wildlife Advisory Group and the ethics committees they represent. Revised previously by Bob Harden (OEH), Mike Fleming, (OEH) and Amanda Paul (DPI).

# data[].attributes.attachment_link.description
Animal Research Review Panel Guideline 10Revised: January 2020 These guidelines have been written to provide guidance for members of Animal Ethics Committees (AECs) and investigators who carry out wildlife surveys, including those for Species Impact Statements or Environmental Impact Statements required under environmental protection legislation (Biodiversity Conservation Act 2016, National Parks and Wildlife Act 1974, Environmental Planning and Assessment Act 1979 and the Threatened Species Conservation Act 1995).

# data[].attributes.body
Link to guidelines:
https://www.animalethics.org.au/policies-and-guidelines/wildlife-research/wildlife-surveys
Primary literature for amphibians:
https://asih.org/sites/default/files/documents/resources/guidelinesherpsresearch2004.pdf

# data[].attributes.body
http://www.environment.gov.au/epbc/publications/survey-guidelines-australias-threatened-frogs

Specific guidelines for doing surveys that involve threatened frogs. This is good as it helps prevent accidental mistreatment to the species being studied,

# data[].attributes.body
Threatened species survey and assessment
guidelines: field survey methods for fauna

https://www.environment.nsw.gov.au/-/media/OEH/Corporate-Site/Documents/Animals-and-plants/Threatened-species/amphibians-field-survey-methods-090213.pdf

# data[].attributes.body
https://www.publish.csiro.au/wr/WR99066

Models of the distribution and abundance of ground-dwelling mammals in the eucalypt forests of north-eastern New South Wales in relation to habitat variables

counts and diversity of populations within different regions based on habitat make up, other variables such as predators also taken into account. Method of counts yet to be determined.

# data[].attributes.headline
General Field Survey methods (threatened species)

# data[].attributes.subject
General Field Survey methods (threatened species)

# data[].attributes.body
https://www.environment.nsw.gov.au/-/media/OEH/Corporate-Site/Documents/Animals-and-plants/Threatened-species/field-survey-method-guidelines.pdf

# data[].attributes.headline
arboreal animals specific to camera trapping

# data[].attributes.subject
arboreal animals specific to camera trapping

# data[].attributes.body
https://link.springer.com/article/10.1007/s13364-012-0122-9

# data[].attributes.body
https://www.environment.gov.au/system/files/resources/b1c6b237-12d9-4071-a26e-ee816caa2b39/files/survey-guidelines-mammals.pdf
Guidelines for detecting mammals listed as threatened species. In
cludes survey design and methods for different species

# data[].attributes.body
https://www.animalethics.org.au/policies-and-guidelines/wildlife-research/wildlife-surveys

# data[].attributes.body
But also contains headings specific to bats, platypus, birds, frogs and reptiles, turtles, and fish.

Australian focused information on general survey techniques, types of traps, ethics, and then more specifics on the animals list above.
Includes lots of link to further reading and has been recently revised (Jan2020)

# data[].attributes.attachment_link.readable_text
Animal Research Review Panel Guideline 10
Revised: January 2020

These guidelines have been written to provide guidance for members of Animal Ethics Committees (AECs) and investigators who carry out wildlife surveys, including those for Species Impact Statements or Environmental Impact Statements required under environmental protection legislation (Biodiversity Conservation Act 2016, National Parks and Wildlife Act 1974, Environmental Planning and Assessment Act 1979 and the Threatened Species Conservation Act 1995).

There are no general texts that describe the ethical capture and handling of all Australian wildlife in the field, although some have been published overseas (see Section 12). There is detailed recent Australian information on the capture, handling and care of native mammals in the National Health and Medical Research Council (NHMRC 2014) A Guide to the care and use of Australian native mammals in research and teaching and this should be used as a primary reference for this group. More general guidance for all animals is available in NHMRC (2013) Australian code for the care and use of animals for scientific purposes, hereafter referred to as the Code. Information on first aid for injured wildlife can be found in Walraven (1990).

These guidelines are not meant to be exhaustive or prescriptive. Their aim is to provide some general principles for reducing the impact on wildlife during surveys, to promote good practice and to raise issues that AECs need to consider when discussing such proposals. The information they contain is taken from the scientific literature and from animal care statements provided by wildlife surveyors.

1 Definitions

Animal Ethics Committee - A committee constituted in accordance with the terms of reference and membership laid down by the Animal Research Act 1985 and the Australian Code for the Care and Use of Animals for Scientific Purposes (NHMRC).

Anabat ™ detector - An electronic device designed to record the echolocation calls of bats (that are usually beyond human hearing) and analyse the characteristic calls of specific species. There are some limitations to this equipment such as inability to distinguish all species. For example it will not detect flying-foxes as they do not use echolocation. Direct observation can be used for these species.

Box trap - A box made from sheet metal with an open door that is released and closes when an animal interferes with the bait in the trap. Sizes vary from quite small (for catching mice) to traps large enough to bandicoots. Because the most commonly used brand is Elliott, these are often referred to as Elliott traps.

Cage trap - Similar to a box trap except that the trap is made from steel mesh. Sizes vary from quite small (for catching mice) to traps large enough to trap dogs. The most commonly used size is 60 x 30 x 30 cm.

Camera trap - A remotely activated camera that is equipped with a motion sensor or an infrared sensor, or uses a light beam as a trigger.

Direct observation - Standing and watching, or walking in a particular direction for certain lengths of time, using binoculars or a spotting scope to detect the range and number of birds or large mammals.

Elliott trap - See box trap.

Fyke net - A net constructed of hoops decreasing in size with webbing between to form a cone shape and one or more funnels inside that prevent trapped fish from swimming out (ie a 'hoop net'), and which also has wings of one or two pieces of netting at the first hoop which are anchored into position with poles. These wings guide the fish into the net. Also known as wing, frame, trap or hoop nets. When used in waterways with platypus, the end of the net must be elevated above water height so that any trapped platypus or turtles do not drown.

Gill net - A net of diamond shaped mesh that is set vertically. The fish are unable to back out because their gill covers get caught in the mesh. When used in waterways with platypus the bottom of the net must be only lighted weighted so the platypus can swim to the surface if caught.

Hair tubes - Small PVC tubes lined with double sided sticky tape with an internal compartment where bait is placed. They may be more efficient and cost effective than the other methods for some rare or trap shy mammals.

Harp trap - An array of thin nylon fishing lines tensioned between two horizontal poles with an escape-proof hessian pocket located below. Bats fly into the lines, fall down undamaged into the pocket and crawl up to roost under a hessian flap where they can roost until released.

Hoop net - See fyke net.

Mist net - Large very fine nylon nets that are strung across potential flyways close to the ground between the vegetation in order to catch birds or bats that fly into them and become entangled. It is very easy for both birds and bats to injure themselves or become distressed whilst being disentangled from these nets so great care is needed.

Playback calls - Pre-recordings of the calls of nocturnal birds (such as owls), frogs and arboreal mammals (such as the koala) which are then played back at night in order to elicit a response from any member of the target species present which may be a reply (or call back) or an approach. They are usually broadcast at various locations over a specified duration (e.g. 10 minutes initial listening, 15 minutes playing of the recording and 10 minutes listening for a response).

Pitfall trap - A glass, metal or plastic container sunk into the ground so that the mouth is level with the soil surface. Ground dwelling animals fall into the trap and are unable to escape.

Proposal - A written outline of a project put forward for consideration by an AEC.

Spider tubes - Small PVC tubes installed into the ground, covered by a metal or canvas roof. Tubes are checked for sheltering individuals which can be captured by hand for identification.

Trip line - A single nylon line stretched 1.5-3cm above the surface of a body of water where bats are likely to fly, causing bats in flight to fall into the water and swim out where they are captured. These have much greater potential for damage to the animal than harp traps.

Voucher specimen - Any specimen that serves as a basis of study and is retained as a reference. A "type" specimen is a particular voucher specimen that serves as a basis for taxonomic description of that subspecies.

Wildlife - Free-living vertebrates of native, non-indigenous and feral species including captive bred animals and those captured from free-living populations.

2 The Regulation of Animal Care during Wildlife Surveys in NSW

In NSW, the welfare of animals used for animal research is protected by the Animal Research Act 1985. Wildlife surveys are considered as animal research and are therefore subject to the provisions of this legislation. The legislation also requires that the conduct of animal research must be consistent with the provisions of the Code.

Wildlife surveyors and AEC members should be conversant with the animal research legislation and the Code, and particularly Clauses 3.3.33 to 3.3.47 of the Code relating to wildlife and field techniques.

In addition to the legislation and the Code, the Animal Research Review Panel (the Panel) has a number of policies and guidelines relevant to wildlife surveys that should be consulted (see Section 13.0). Investigators should be familiar with the contents of all these documents. Along with general information on the legislation, AECs, the Panel and the Code, the policies and guidelines are available on the Animal Ethics Infolink site.

The intent of the legislation is to protect the welfare of animals and is implemented through the requirement that all animal research must be approved by an AEC before the work commences. The AEC considers the ethical aspects of the study such as the justification for the survey, the methods used and the impact of all procedures on the animals. To assist the AEC in its deliberations, application is made by the investigator on a proposal form that aims to ensure that AEC members have the information they need for their discussions, written in plain English.

Intending wildlife surveyors associated with an Accredited Animal Research Establishment should seek approval from the establishment's AEC. Those not associated with an Accredited Establishment may apply to the Secretary's AEC through the Department of Primary Industries. If the AEC approves the application, the investigator will be issued with an Animal Research Authority for the project.

A Biodiversity Conservation Licence (also referred to as a Scientific Class Licence) issued under Part 2, Division 3, of the Biodiversity Conservation Act 2016 to conduct research is also required from the Office of Environment and Heritage to conduct wildlife surveys in NSW. This licence requires a current Animal Research Authority.

3 General ethical considerations and wildlife surveys

the replacement of animals with other methods;

the reduction of the number of animals used; and

the refinement of the techniques used to reduce the impact on animals.

The Code advances three fundamental concepts for improving the welfare of animals used for scientific research. These are known as the Three Rs and include:

1. Survey planning
Replacement of animals may be possible by assessing the existing recent knowledge of the fauna in the proposed study area and determine if a new survey is both necessary and justified. Sources of such information include:

published records in scientific journals and the newsletters of scientific and natural history societies. If a survey has been done many years ago in the same area a current survey is probably warranted to allow an assessment of possible changes in species abundance;

biodiversity survey reports by the Office of Environment and Heritage, including NSW National Parks and Wildlife Service and Forestry Corporation of NSW;

the Atlas of NSW Wildlife - a whole-of-government system for flora and fauna sightings information;

Birdlife Australia The New Atlas of Australian Birds (Barrett et al. 2004);

Atlas of Living Australia;

previous Species Impact Statements and Environmental Impact Statements available from the Department of Planning and Environment; and

local knowledge (local councils, land management groups, land holders).

Historical records may not be useful in determining current species presence or abundance as these may have changed over time, often due to threatening processes. In cases such as these surveys may have particular value as they may show trends in species distribution and abundance over time.

Reduction in the usage of animals may be possible if the background research suggests recent surveys have been undertaken in an area of interest and therefore a reduction in survey effort may be appropriate. Wildlife surveyors are encouraged to publish survey information whenever possible, and to lodge results where they can be accessed in the future. Examples of such repositories include the Australian Museum and the NSW Wildlife Atlas and the Atlas of Living Australia.

Refinement in survey techniques comes from substituting non-invasive techniques such as camera traps where they can provide the same information as conventional trapping techniques.

2. Survey design and methodology
It is not the purpose of these guidelines to provide detailed technical advice about the design and methodology of wildlife surveys. Rather, the intent is to show how the welfare of animals can be protected during surveys by employing appropriate design and methodology. This section is a general discussion of these issues. In later sections more specific suggestions are made.

The following points should be considered when designing a wildlife survey:

The methods used should be appropriate to the objectives of the project;

The methods used should be based on sound scientific principles so that the results are valid;

The methods used should minimise the impact on the animals; and

Sample sizes should be kept to the minimum required statistically and that number should be able to be justified.

The following general points should be considered when determining the methods to be used in a wildlife survey.

3. Voucher specimens
The collection of voucher specimens is an important part of scientific research. However, it is a practice of concern to some sections of the community. Wildlife surveyors intending to collect voucher specimens should consult the Panel's guidelines for the Collection of voucher specimens. With the recent advances in genetic screening, consider taking only a tissue sample that can be used for genetic analysis. Consult on an appropriate method with a genetic repository such as the Australian Museum or for mammals refer to NHMRC (2014).

Some important points are listed below:

4. Emergency procedures
All applications to an AEC for wildlife research require a detailed description of emergency procedures in the event of injuries to animals, inclement weather, floods, bushfires and the illness or injury of the surveyors. The purpose of these is to ensure that threats to the welfare of animals resulting from emergencies are identified and mitigated.

Investigators should be conversant with the Panel's Emergency Procedures policy. Issues particularly relevant to wildlife surveys include the following.

Arrangements must be made to clear and close all traps in the event of inclement weather, floods and bushfires.

Arrangements must be made to clear and close traps in the event that illness or injury removes the investigator from the field.

Investigators should have the appropriate skills and equipment to euthanase seriously injured animals in the field should this be necessary. Euthanasia must be by an approved method (see section 3.5 below).

Arrangements must be made to appropriately transport seriously injured animals to the nearest veterinarian for treatment, noting that injured animals should be taken to veterinarians initially rather than to wildlife carers.

Any unexpected problems should be reported to the AEC as soon as possible, including mortalities and injuries to animals. Future surveys may need to be modified in the light of these problems.

5. Euthanasia
Emergency euthanasia or killing of specimens for vouchers may need to be carried out by surveyors in the field. The following points about euthanasia need to be considered.

Methods must be humane and produce a painless death as rapidly as possible.

Methods that are acceptable are described in UFAW/WSPA (1989), UFAW (1996, 1997), Reilly (2001) and AVMA (2013). For native mammals the methods referred to in NHMRC (2014) can be followed.

Methods that are not acceptable include car exhaust fumes, cervical dislocation in animals larger than 150g, drowning and freezing. Note that cooling reptiles and amphibians to make them easier to handle is acceptable but, even after cooling, freezing is not an acceptable method of euthanasia.

Surveyors must be trained and competent in the use of the acceptable methods of euthanasia.

Investigators should be conversant with the Panel's policy, The use of restricted drugs and the conduct of restricted acts of veterinary science in animal research, which details the legal requirements for using sodium pentobarbitone.

6. Diseases and parasites

The Three Rs are as relevant to wildlife surveys as to laboratory studies of animals, and in this section examples of their implementation through good survey planning and design and appropriate methodology are discussed. Specific wildlife survey methods are discussed in later sections.

Surveyors should be aware of their potential exposure to zoonotic diseases (those that affect both animals and humans and may be passed between them) that are present in Australian native animals. Some of these diseases can cause serious illness or death. Several examples of zoonotic diseases include:

Australian Bat Lyssavirus - a virus carried by many species of bats that usually causes death unless vaccinated against.

Psittacosis - caused by the bacterium Chlamydia psittaci that is typically carried by birds, especially pigeons and parrots.

Q fever - Caused by exposure to the bacterium Coxiella burnetii that is carried by livestock and wild life.

Salmonellosis - Caused by exposure to the bacterium Salmonella that is carried by reptiles and various other wildlife.

Toxoplasmosis - Caused by the protozoan Toxoplasma gondii that is carried by cats.

Viral and bacterial infections:

mites and ticks - These are known to be carries or various diseases including Australian Tick Typhus, Lyme disease, Babesiosis, tick paralysis and Red Meat Allergy. These infect people when they bite.

External parasites:

cestodes (tapeworms).

nematodes (roundworms).

trematodes (flukes).

Internal parasites:

high levels of personal hygiene;

not eating, drinking or smoking whilst handling animals;

washing field clothes and equipment that has come into contact with animals' blood or body fluids and cleaning all survey equipment between surveys;

basic first aid for treatment of cuts, bites and scratches;

observance of protocols to avoid transmission of frog chytrid fungus as outlined (Department of Environment and Climate Change 2008);

obtaining vaccinations against Australian Bat Lyssavirus before handling microbats and/or flying foxes.

minimise exposure to ticks.

Surveyors should therefore take basic precautions to prevent animal-animal, animal-human and human-animal transfer of disease. Such precautions include the following:

Should anyone who handled animals become ill within two months of a survey, the attending medical practitioner should be informed of the potential exposure to zoonoses. Further information on zoonoses can be obtained from NSW Health.

4 Surveys of terrestrial and arboreal animals.

The Environment, Energy and Science Group has published a number of guidelines on biodiversity surveys and how to sample native fauna. There are also specific guidelines on how to survey for NSW threatened species.

1. Methods not involving animal capture
Some mammal species leave signs (scats and tracks) sufficiently distinctive to provide positive identification. The sign of many Australian mammal species is described in Triggs (1996). Signs that indicate the presence of species or groups of species should be used in surveys wherever possible.

2. Animal signs

3. Hair tubes
The use of hair tubes is described by Scotts and Craig (1988), Lindenmayer et al. (1999) and Mills et al. (2002). Points to consider are:

Ensure that the floor of the tube is free of adhesive tape to prevent small lizards and frogs becoming stuck.

If an animal does become stuck to the tape, do not try to pull the tape off, as this may seriously damage the skin. Either carefully trim the tape on the animal to as small a size as possible (the remaining tape will be shed during normal skin replacement) or gently ease vegetable oil under the tape and slide it off.

Slope hair tubes with the entrance pointing slightly downwards to ensure drainage.

Commercially designed hair funnel traps are available with a specially prepared sticky wafer that minimise accidental captures of animals.

4. Spotlight counts
When spotlighting animals:

avoid prolonged exposure to the light (i.e. more than two minutes);

use a light with a narrow beam; and

when practical, use a red filter or, preferably, a dimmer switch to reduce light intensity for prolonged observations once the animal has been spotted.

Survey with a low wattage light (30 to 50 w) and briefly use a narrow beam of brighter light for identification (SEWPAC 2011a).

5. Camera traps

The Commonwealth Department of the Environment has also prepared survey guidelines for nationally listed mammals, bats, birds, reptiles, frogs and identify techniques are applicable to the species level. See SEWPAC (2010a, 2010b, 2011a, 2011b).

Traps may be infra-red or white flash.

Infra-red cameras are less intrusive but do not distinguish some species sufficiently, as they do not distinguish colour.

Use of a lure, either scent or mechanical, focuses the detection zone and holds the animal long enough to get an image.

Often used in conjunction with other survey methods.

1. Methods involving animal capture

2. Trapping - general

Use the trapping method with the least impact.

Whenever possible, avoid trapping at times of the year when animals may be susceptible to greater stress, such as during breeding seasons or extreme heat or cold. If animals are breeding, minimise their time in traps by checking more frequently and releasing pregnant or lactating females as a matter of priority.

Select the type of trap that is appropriate to the species being targeted.

Ensure all traps are in good working order, cleaned and checked immediately prior to use.

Limit the number of traps set per field worker to that which can be cleared in two hours.

At any one site, unless otherwise justified, limit trapping periods to no more than four consecutive nights with a minimum of three nights between trapping periods to avoid continually trapping the same individuals.

Use a bait appropriate to diet of the target species. The bait should not only lure the animal into the trap, but should also replace the food and moisture it would have consumed had it not been trapped. This is particularly important for small mammals that have high metabolic rates.

Locate each trap to reduce exposure of trapped animals to the sun, wind, rain etc (for example, place traps under shrubs or beside logs).

Avoid placing traps in areas of high ant activity.

Do not trap during periods of badly inclement weather.

Ensure all traps are located and checked each time a trapline is checked and that all traps are removed from the field or closed at the end of the trapping period. If individual traps are numbered and set in order, it makes it easier to ensure that all traps are checked.

For nocturnal species, begin clearing traps at first light and where practical leave the traps closed until late afternoon. During periods of extremely cold weather, cease trapping completely or clear and close traps by 0200 hrs each day.

For diurnal species, have an inspection schedule that minimises the impact on any trapped animals and locate the traps so as to minimise the possibilities of heat or cold stress.

Release animals as soon as possible and where they were caught.

Cease trapping immediately if there has been an unusually high mortality of animals.

1. Box traps (also known as Elliott traps)
In addition to the general points in 4.2.1 above, the following need to be considered.

Provide bedding in the traps. Dry leaf litter and Dupont Hollofill ™ are suitable materials, although the latter sometimes wraps around the animals' feet. Cotton wool should not be used because it absorbs moisture, increasing the risk of hypothermia.

In areas with wetter climates, place traps in a plastic bag, taking care to ensure adequate drainage (slope traps at 10o to the horizontal to allow drainage during rain).

During periods of high temperatures in areas where traps cannot be sheltered from the sun, close traps during the day.

Traps set in trees should be on the opposite side of the tree to the morning sun.

2. Cage traps
In addition to the general points in 4.2.1 above, the following need to be considered.

Set traps in sheltered positions.

Provide shelter for trapped animals by covering the trap with opaque plastic or canvas (cooler areas) or with hessian or shade cloth (hotter areas).

If traps cannot be sheltered from the sun, they should be closed during the day if temperatures are high.

3. Dry pitfall traps
The Panel has developed guidelines for the use of pitfall traps, which should be observed. In addition, consider the following points:

4. Radio-tracking

The use of camera traps has become widespread in fauna survey and applies not only to mammals but also birds and reptiles (SEWPAC 2011a, Meek et al. 2012). Points to consider are:

In general, the following points apply to the use of traps.

1. Methods not involving animal capture
Ultrasound detectors (for example, the AnaBat ™, Song Meter ™) can be used to detect bats without any impact and should be used whenever possible.

2. Methods involving animal capture

3. General

Radio transmitters are rarely necessary for general wildlife surveys so their use is not covered here in depth. See the Panel guideline Radio tracking and GPS tracking.

Whenever possible avoid trapping during the breeding season.

Bats should be released at the point of capture as soon as possible. However, they should not be released in daylight. Those that cannot be released before dawn should be held until the following dusk.

When necessary, bats should be held separately in suspended cloth bags in a dark, quiet and warm place.

Bats may go into torpor in the trap or while held in bags and will need to be re-warmed before release.

Care should be taken when handling both flying foxes and microbats, due to the zoonotic disease Australian bat lyssavirus (see Section 3.6)

1. Harp traps
A description of the use of harp traps can be found in Tidemann and Woodside (1978). Points additional to those in 5.2.1 that need consideration are:

set traps in a sheltered spot in potential flyways;

clear within two hours of dusk and again after dawn but before the sun begins to warm the hessian; and

harp traps must not be used where large numbers of bats could be caught (for example at entrances to roost sites) to avoid the overheating of bats in the collection bag.

2. Mist nets
Points additional to those in 5.2.1 that need consideration are listed below.

3. Trip lines

5 Surveys of bats

Risks of injury to bats, so use other methods whenever possible;

Monitor continually whenever the line is deployed;

Be prepared to enter the water to rescue bats if necessary; and

Have at least one low-powered torch to collect bats since they will swim away from bright lights.

A description of bat survey methods can be found in SEWPAC (2010a). Surveys for bats should be carried out by an experienced bat investigator as (apart from the fruit bats) little is known of their biology or taxonomy and species can be difficult to identify.

The following general points need to be considered when trapping bats:

Before any trapping is undertaken, the likelihood of platypus being present should be ascertained from other sources including recent published reports and observational surveys undertaken by the surveyor.

In deep water habitats, lightly weighted gill nets should be used in case platypus are present to allow them to swim to the surface.

Trapping must not be carried out during the months of October-March, when females are laying eggs and raising young, unless justified to the AEC.

Gill nets should be observed continuously to detect when a platypus is caught, e.g. by the splashing.

Gill nets should be lifted every hour to check for snags and entrapped large fish.

Nets should have a stretched mesh size of at least 150 mm, or be fitted with an entrance grill with a mesh size of less than 35 mm, or be set so that there is an air space available along the length of the net.

Due to the potential to cause deaths of platypus and turtles check with NSW Fisheries before deployment. Note that it is illegal to use opera-house style traps in some NSW waterways.

Captured platypus should be transferred to a clean bag or secure box and held in a quiet place until all nets are closed for the night, to minimise the likelihood of hypothermia, attack by predators or by other platypus (particularly between males in the breeding season).

Note that trip lines are ineffective for bats that can take off from water (e.g. fishing bats).

Points additional to those in 5.2.1 that need consideration are:

1. Methods not involving animal capture

Avoid close range inspection during breeding and feeding.

Carry out searches for nests, mounds, display areas, characteristic scrapes and scratchings, visual and auditory searches such as breeding calls.

2. Direct Observation

3. Playback calls

Avoid prolonged exposure by limiting calling sessions to two 15 minute periods per night.

Use of play back calls during the species' breeding season should be done with care so as not to disrupt the breeding of the resident animals.

4. Acoustic Monitoring
The advent of cheap digital acoustic recorders that allow the continuous recording of ambient noise has allowed acoustic monitoring to become a viable method of survey. Subsequent sound analysis, using specialised software or expertise, identifies the calls of particular species. This could be an efficient method of detecting species that only occasionally frequent an area or are highly secretive and difficult to detect.

5. Spotlighting owls

6 Platypus

Platypuses are one of the most difficult species to handle and maintain in captivity, because they are highly susceptible to stress unless managed carefully and with a high degree of knowledge and expertise (NHMRC 2014). Wildlife surveys would rarely need to capture platypus to establish their presence. Direct observation at dawn and dusk of suitable riverine habitat is sufficient to establish their presence but not their absence.

7 Surveys of birds

Because of the high risk of injury and death to birds, mist nets should only be used where other methods have already been rejected as unsuitable.

Mist nets must only be used by trained and competent personnel. An appropriate authority for the use of mist nets by the Australian Bird and Bat Banding Scheme (ABBBS) is preferred.

Mist nets should be attended at least every 30 minutes and captured birds removed immediately.

Nets must be closed when not attended.

Numerous methods have been developed for the detection of birds in wildlife survey and the structuring of sample design (SEWPAC 2010b). Choosing an appropriate design and method will depend entirely on the questions being asked.

1. General
A summary of survey methods for reptiles can be found in Blomberg and Shine (1996).

Choosing the correct season is critical for effective surveys of frogs (and to a lesser extent with reptiles). Most frog species are active only during the warmer months of the year (Spring-Summer-Autumn), although there are some that are active only during the cooler months (Autumn and Winter). Outside of their active season many frogs aestivate or go into torpor, usually in burrows, hollows in trees, crevices in timber or rocks or under loose soil. When in torpor, they are undetectable. To a lesser extent this may also occur during the active season when weather conditions are unsuitable (e.g. dry).

2. Surveys not involving animal capture

3. Spotlighting frogs with or without using playback

Avoid excessive foot traffic around the water body.

Keep exposure to a minimum to prevent overheating.

Use a lower intensity light held at a distance for further observations.

1. Camera Traps
Camera traps can be deployed for reptiles when mounted vertically over a gap in a drift fence. This works particularly well for larger lizards and snakes that are not readily captured with conventional means. See Welbourne (2013, 2014) for this technique.

2. Surveys involving animal capture

3. General

1. Freshwater turtles

Set traps with an air space to prevent drowning of turtles or by-catch such as platypus, water rats or water birds. The air space can be maintained by use of a float (e.g. an empty drink container) or by tying the trap to an overhanging tree or log. Opera-house style traps can be tied to a stake on the bank. Note that is illegal to use opera-house style traps in some NSW waterways; check with NSW Fisheries before deployment.

Traps should be checked at least at dawn and dusk. They should be checked more frequently if turtle numbers are high and during summer.

Transport animals separately to avoid the risk of shell damage and hence infection. Keep cool during transport to avoid heat stress.

2. Marine turtles

Marine turtles are very susceptible to heat stress, especially during transport. They can be cooled by the use of wet hessian bags.

Confining the animals in small spaces increases the risk of abrasions, and hence infections. Marine turtles are best restrained by placing them on their backs in a cool place.

During transport, insulate from heat and also from vibration. They are best transported within a vehicle rather than in the tray of a utility.

Consider that hand searches carried out by experienced personnel under suitable conditions will locate nearly all species of reptiles and frogs in an area within a short period of time that may mean that fewer traps or no traps at all need be set.

Frogs should be handled as little as possible because handling removes skin secretions and predisposes the frog to fungal infections (White 1990), while continuous holding in the hand can result in overheating.

Hygiene precautions as detailed in Department of Environment and Climate Change (NSW) (2008) must be observed when handling frogs and tadpoles, including the use of gloves.

Gloved hands should be made wet in the local water or in wet grass/vegetation so that loss of skin secretions is minimised when frogs are first picked up.

Frogs should be moistened with rainwater or water from the stream being surveyed after holding. They may be held separately temporarily (up to 24 hours) in a new moist plastic bag containing some vegetation (although, in the dark, vegetation will absorb oxygen).

Reptiles should be held separately in appropriately sized secure bags or boxes with some vegetation, or a moist paper towel, as appropriate, in a cool place.

Tadpoles are often easier to find than adult frogs and provide important information about habitats used and other measures of environmental quality. However, care needs to be taken when handling tadpoles, as handling can result in a high level of injury and death of the tadpoles. See Anstis (2002) for identification keys.

Consider that fish are usually in their best condition in spring and early summer and will be able to cope with the shock of capture and recover more quickly than in the winter or in mid-summer after spawning.

Use nets with soft mesh (for example, cotton or nylon) to reduce damage to the fish.

Use appropriately sized and weighted traps to reduce the risk of non-target animals being caught.

Fyke nets should have an air space by being set partially out of the water to prevent drowning of trapped mammals (such as platypus and water rats) or waterfowl. Otherwise they should have a means of escape.

If possible, avoid using gill nets because fish caught in these often die (or are so damaged during removal that they are unlikely to survive) and because they have the potential to trap many non target species.

Check and empty traps regularly.

Handle the fish as little as possible.

Minimise the removal of the fish's protective mucous covering and reduce temperature shock by wetting hands first in the water that the fish was caught.

If electrofishing is being used for sampling, operators should have appropriate training and follow guidelines set out in NSW Fisheries (1997).

1. Hand searching for amphibians and reptiles

Take care when moving and repositioning rocks and logs to prevent animal injuries and avoid causing habitat disturbance that may affect the subsequent abundance of the species.

Wash hands without soap (for instance in the water of the water body being surveyed or with rainwater) to reduce contamination from chemicals.

Noose type devices to catch large reptiles should be used with care and devices used to pin snakes need to be padded to avoid causing injury.

2. Pitfall traps
Note that the Panel's guideline Use of pitfall traps should be followed.

See also section 4.2.4 Dry pitfall traps.

3. Spider tubes

Examples of techniques to census owls can be found in Kavanagh and Peake (1993).

See also section 4.1.3 Spotlight counts

Guidelines for using mist nets can be found in the Australian Bird Banders Manual (Lowe 1989).

8 Surveys of reptiles and frogs

Some species of reptile including Grassland Earless Dragon inhabit burrowing spider tubes. Inspection of these visually or with an endoscope can be used to detect the presence of these species.

9 Surveys of turtles

10 Surveys of fish
Department of Environment and Climate Change (NSW) (2008) Hygiene protocol for the control of disease in frogs. Information Circular No. 6. DECC (NSW). Sydney South
General information on survey design and best use of methods is available in Merrick (1990), SEWPAC (2011b) and Barker et al. (2009).
Fair, J., Paul, E. & Jones, J. (Eds) (2010) Guidelines to the use of wild birds in research. Ornithological Council, Washington DC.
11 References

American Society of Ichthyologists and Herpetologists (2004) Guidelines for the use of live amphibians and reptiles in field and laboratory research.Second Edition. Herpetologists' League, Society for the Study of Amphibians and Reptiles. 42pp.

American Veterinary Medical Association (2013) AVMA Guidelines for the euthanasia of animals: 2013 edition. American Veterinary Medical Association, Schaumburg, Illinois.

Anstis, M. (2002) Tadpoles of South-east Australia. New Holland Publishers, Sydney.
Meek, P. & Fleming, P. (eds.) (2014) Camera Trapping: Wildlife Management and Research. CSIRO Publishing, Melbourne.
Bali, R. & Delaney, R. (1996) Assessment of Koala Radiocollaring Studies in South eastern Australia. NSW National Parks and Wildlife Service, Sydney.
National Health and Medical Research Council (2013) Australian code for the care and use of animals for scientific purposes, 8th edition. Canberra: National Health and Medical Research Council. National Health and Medical Research Council (2014) A Guide to the care and use of Australian native mammals in research and teaching. National Health and Medical Research Council, Canberra. NSW Fisheries (1997) Australian Code of Electrofishing Practice. NSW Fisheries Management Publication No. 1
Barker, D., Allan, G.L., Rowland, S.J., Kennedy, J.D. & Pickles, J.M. (2009) A Guide to Acceptable Procedures and Practices for Aquaculture and Fisheries Research. Primary Industries (Fisheries) Animal Care and Ethics Committee, Nelson Bay NSW.

Barrett, G., Barry, S., Cunningham, R., Poulter, R. & Silcocks, A. (2004) The New Atlas of Australian Birds. CSIRO Publishing, Melbourne.

Blomberg, S. & Shine, R. (2006) Reptiles, pp. 297-307, in J. Sutherland (ed.) Ecological Census Techniques: A Handbook. Second Edition. Cambridge University Press, Cambridge.

DeTolla, L.J., Srinivas, S., Whitaker, B.R., Andrews, C., Hecker, B., Kane, AS. & Reimschuessel, R. (1995) Guidelines for the Care and Use of Fish in Research. ILARJournal 37: 159-173.

Helman, P. & Churchill, S. (1986) Bat capture techniques and their use in surveys. Macroderma 2: 32-53.

Kavanagh, R. & Peake, P. (1993) Survey procedures for nocturnal forest birds: an evaluation of the variability in census results due to temporal factors, weather and technique. Pp. 86-100, in P. Olsen (ed.) Australian Raptor Studies. Australian Raptor Association, RAOU, Melbourne.
Sikes, R.S. & Gannon, W.L. and the Animal Care and Use Committee of the American Society of Mammalogists. (2011) Guidelines of the American Society of Mammalogists for the use of wild mammals in research. Journal of Mammalogy 92: 235-253. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2010a) Survey guidelines for Australia's threatened bats. Commonwealth of Australia, Canberra. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2010b) Survey guidelines for Australia's threatened birds. Commonwealth of Australia, Canberra. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2011a) Survey guidelines for Australia's threatened mammals. Commonwealth of Australia, Canberra. Sustainability, Environment, Water, Population and Communities (SEWPAC) Department (2011b) Survey guidelines for Australia's threatened fish. Commonwealth of Australia, Canberra.
Lindenmayer, D.B., Incoll, R.D., Cunningham, R.B., Pope, M.L., Donnelly, C.F., MacGregor, C.I., Tribolet, C. & Triggs, B.E. (1999) Comparison of hairtube types for the detection of mammals. Wildlife Research 26: 745-753.

Lowe, K.W. (1989) The Australian Bird and Bat Banders Manual. Australian National Parks and Wildlife Service, Canberra.

Mills, D.J., Harris, B., Claridge, A.W. & Barry, S.C. (2002) Efficacy of hair-sampling techniques for the detection of medium-sized terrestrial mammals. I. A comparison between hair-funnels, hair tubes and indirect signs. Wildlife Research 29: 379-387

Parnaby, H. (1992a) An interim guide to the identification on insectivorous bats of southeastern Australia. Technical Reports of the Australian Museum No. 8. Australian Museum, Sydney.

Parnaby, H. (1992b) Summary of the ultrasonic survey of the microbats of north east New

South Wales, 1991/1992. Report to the NSW National Parks and Wildlife Service, Sydney.

Petit, S. & Waudby, H.P. (2012) Standard operating procedures for aluminium box, wire cage, and pitfall trapping, handling, and temporary housing of small wild rodents and marsupials. Wildlife Research 60: 392-401.

Reilly, J. (ed) (2001) Euthanasia of animals used for scientific purposes. ANZCCART, Glen Osmond, South Australia.

Scotts, D.J. & Craig, S.A. (1988) Improved hair sampling tube for detection of rare mammals. Australian Wildlife Research 15: 469-472.

Tidemann, C.R. & Woodside, D.P. (1978) A collapsible bat trap and a comparison of results obtained with the trap and mist nets. Australian Wildlife Research 5: 355-362.

Triggs, B. (1996) Tracks Scats and Other Traces: A Field Guide to Australian Mammals. Oxford University Press, Melbourne.

Universities Federation for Animal Welfare (1996) Recommendations for euthanasia of experimental animals: Part 1. Laboratory Animals 30: 293-316.

Universities Federation for Animal Welfare (1997) Recommendations for euthanasia of experimental animals: Part 2. Laboratory Animals 31: 1-32.

Universities Federation for Animal Welfare /World Society for the Protection of Animals (1989) Euthanasia of Amphibians and Reptiles: Report of a Joint UFAW/WSPA Working Party. UFAW, South Mimms, Potters Bar, Herts, UK. And WSPA, London, UK.

Van Dyck, S. & Strahan, R. (2008) The Mammals of Australia. New Holland Publishers, Sydney.

Walraven, E. (1990) Taronga Zoo's Guide to the Care of Urban Wildlife. Allen and Unwin, Sydney.

Welbourne, D. (2013) A method for surveying diurnal terrestrial reptiles with passive infrared automatically triggered cameras. Herpetological Review 44: 247-250.

Welbourne, D.J. (2014) Using camera traps to survey diurnal terrestrial reptiles: a proof of concept. pp. 225-232, P. Meek, P. Fleming, G. Ballard, P. Banks, A. Claridge, J. Sanderson & D. Swann D (eds.) Camera Trapping Wildlife Management and Research. CSIRO Publishing, Melbourne.

Acknowledgments

Stephen Jackson (NSW DPI) for the development of the current edition and Mike Fleming (DPIE) for providing comments on the draft document. The previous edition of these guidelines was developed with the assistance of M. Murray, P. Burcher, A. Adair, K. Kendall, Leong Lim, ERM Mitchell McCotter, Bob Harden, Gerard Körtner, Fritz Geiser the Australian Platypus Conservancy, the Victorian Department of Natural Resources and Environment, members of the Wildlife Advisory Group and the ethics committees they represent. Revised previously by Bob Harden (OEH), Mike Fleming, (OEH) and Amanda Paul (DPI).

# data[].attributes.attachment_link.description
Animal Research Review Panel Guideline 10Revised: January 2020 These guidelines have been written to provide guidance for members of Animal Ethics Committees (AECs) and investigators who carry out wildlife surveys, including those for Species Impact Statements or Environmental Impact Statements required under environmental protection legislation (Biodiversity Conservation Act 2016, National Parks and Wildlife Act 1974, Environmental Planning and Assessment Act 1979 and the Threatened Species Conservation Act 1995).

# data[].attributes.body
C.L Hourigan, et. al., 2008, 'A comparison of the effectiveness of bat detectors and hap traps for surveying bats in an urban landscape.

# data[].attributes.body
https://www.nps.gov/caco/learn/nature/upload/CACOwaterbird_monitoring_protocol_jan-2023_03.pdf
" A Protocol for the Long-term Coastal Ecosystem Monitoring Program at Cape Cod National Seashore"
Probably a bit overkill, and may not be applicable to Australia in all regards, but on the other hand very detailed and specific. Can be useful as an overview or when looking at a specific technique.

# data[].attributes.body
https://www.researchgate.net/publication/201999856_Designing_a_Cost-Effective_Invertebrate_Survey_A_Test_of_Methods_for_Rapid_Assessment_of_Biodiversity

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Specific to bats: Species credit’ threatened bats and their habitats: NSW survey guide for the Biodiversity Assessment Method

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Specific to bats: Species credit’ threatened bats and their habitats: NSW survey guide for the Biodiversity Assessment Method

# data[].attributes.body
https://www.environment.nsw.gov.au/-/media/OEH/Corporate-Site/Documents/Animals-and-plants/Threatened-species/species-credit-threatened-bats-survey-guide-180466.pdf

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https://www.publish.csiro.au/wr/wr04094

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GUIDELINES
https://www.environment.gov.au/system/files/resources/107052eb-2041-45b9-9296-b5f514493ae0/files/survey-guidelines-birds-april-2017.pdf

2008 Aus government document on Survey guidelines for Australia's threatened birds.
Contains specific guidelines for different species.

PAPER
https://wildlife.onlinelibrary.wiley.com/doi/epdf/10.1002/jwmg.821?saml_referrer
2015 paper that uses automated recorders to monitor forest birds across Northern California. This information was needed to inform conservation planning efforts.
Useful as it outlines the survey methods used and the species observed.

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https://www.environment.gov.au/epbc/publications/survey-guidelines-australias-threatened-birds

Guidelines for presence/absence surveys as per the federal government. Therefore, useful for this taxa and this method of survey.

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Existing Guideline (General) (https://birdata.birdlife.org.au/survey-techniques)

Systematic Bird Surveys

2-ha, 20 min Search

Area Search (500m/5km)

Embedded Surveys (Two surveys in one)

2-ha, 20 min Search

This involves searching for birds in a two-hectare area for 20 minutes. The recommended shape for the two hectares is 100 m x 200 metres. You can use other shapes, such as a circle with a radius of 80 metres, or a strip 400 m long x 50 m wide. Only record birds within the two-hectare area (though see Embedded Survey below). Birds flying over the search area should be included.

Primary Literature 1: (https://www.sciencedirect.com/science/article/abs/pii/S1574954117302467

Bird species identification via transfer learning from music genres

This paper proposes to transfer knowledge existing in music genre classification to identify bird species, motivated by the existing acoustic similarities.

# data[].attributes.body
http://www.environment.gov.au/system/files/resources/2f420bf1-d9e4-44ec-a69c-07316cb81086/files/survey-guidelines-bats.pdf

- survey guidelines for Australia's threatened bats
- its 72 pages, i think its going to be very useful haha, also its a government document.

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https://www.publish.csiro.au/wr/wr00041
general population counts following post fire changes to environment- useful as it is after abundance and diversity counts

# data[].attributes.body
https://www.researchgate.net/publication/248883441_A_Comparison_of_Techniques_Used_in_a_Survey_of_the_Ground-dwelling_and_Arboreal_Mammals_in_Forests_in_North-eastern_New_South_Wales

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https://www.environment.nsw.gov.au/-/media/OEH/Corporate-Site/Documents/Animals-and-plants/Conservation-management-notes/watching-surveying-wildlife-conservation-management-notes-110655.pdf

4 page summary. Includes nocturnal animals too.  Includes multiple passive techniques, real time observation and inference observation (animal tracks, feathers, faeces, etc). Contains references to other resources. Good starting point to refine search.

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Specific to Amphibians
Comparison of three different techniques in an area in Washington.
https://geo.nwifc.org/CMER/PublicDocs/CMER_TOC/Documents/Table_of_Contents/CMER_Projects/SAASensitiveSitesIDMeth/TOC_F5_final%20seep%20study%20.pdf

Specific to Amphibians
Comparison of 3 techniques in SE Queensland
https://www.jstor.org/stable/3893088

Specific to Amphibians
Govt guidelines for field survey methods for threatened amphibians- explanation of many techniques
https://www.environment.nsw.gov.au/research-and-publications/publications-search/threatened-species-field-survey-methods-for-fauna-amphibians

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https://www.environment.nsw.gov.au/licences-and-permits/wildlife-licences/catch-and-release-licence/reptile-catch-and-release-licences

Explains the laws in regard to catching and releasing reptiles and it is useful as it provides an insight into the legal requirements of the task.

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Scientific Licenses NSW Department of Planning, Industry & Environment

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Scientific Licenses NSW Department of Planning, Industry & Environment

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https://www.environment.nsw.gov.au/licences-and-permits/scientific-licences

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guidlines for surveying reptiles in aus