Biological Sciences, Santa Barbara City College pipeline center for sustainability

Biology 130: Methods in Field Biology

Field Technique:  Mammal Trapping



(From Gannon et. al. 2007 with some editting)

Common reasons to capture mammals include livetrapping to tag (with radiotransmitters, necklaces, ear tags, or passive integrated transponder [PIT] tags), mark (number, band, hair color, freeze brand, ear tag, or toe clip), or tissue collection. Regardless of the approach, potential for pain, distress, or suffering must be considered. When livetrapping, adequate insulation, food, and avoidance of temperature extremes must be provided.

 

woodrat release
(Woodrat being released from a Sherman Live Trap. Photo by Adam Green)

 

Live Capture Investigators conducting research requiring live capture of mammals assume the responsibility for using humane methods that respect target and nontarget species in the habitats involved. Methods for live capture include those designed for small mammals (Sherman, Tomahawk,and Hav-A-Hart traps, pitfalls, artificial burrows, and nest boxes), medium-sized to large mammals (Tomahawk, Hav-A-Hart, and foot-hold traps, snares, corrals, cannon nets, culvert traps, and darting), bats (mist nets, harp traps, and bags), and fossorial mammals (e.g., Baker and Williams 1972; Hart 1973). Methods of live capture should not injure or cause excessive stress to the animal. Adequate measures should be taken to ensure that the animal is protected from predation and temperature extremes and has food and water available, as needed, until it is released. For permanent trapping grids or webs, the investigator might provide shelters over traps to protect captured animals from extreme temperatures and precipitation (Kaufman and Kaufman 1989). Use of steel foot-hold traps for capturing animals alive must be approached cautiously because of potential for injury and capture of nontarget species (Kuehn et al. 1986). For some taxa, foot-hold traps, including leg snares, might present the only means of capture available and indeed might be most effective (Schmintz 2005; see also http://www.furbearermgmt. org/resources.asp#bmps for specific techniques). When their use is appropriate, investigators have an ethical obligation to employ steel foot-hold traps of a sufficient size and strength that the animal is held firmly. Traps other than snares with rubber padded or offset jaws should be used to minimize potential damage to bone and soft tissue. Snares or spring foothold traps must be checked frequently (perhaps twice daily or more often depending upon target species and potential for capture of nontarget species) and captured animals assessed carefully for injury and euthanized when necessary. Nontarget species, if uninjured, should be released immediately.

 

Trap and flagging

 

To facilitate prompt checking, the number of traps set at a particular time and location should not exceed the ability of the investigator to monitor traps at reasonable intervals. Because the most effective way to prevent mortality or injury to animals in live traps is prompt and frequent checks of traps for captured animals, the investigator should consider staking or visibly flagging a trap line (or otherwise devising a system) to ensure that all traps are checked and removed reliably and efficiently.

(Orange flagging tape used to mark location of trap. In this set the traps are also numbered and placed about 20 paces apart. This way the researchers know if they skip one because they will miss a number and end up walking double the ditance to the next trap. Photo by Adam Green)

 

Regular monitoring ensures that target animals remain in good condition while in traps and allows prompt release of nontarget species with no ill effects caused by capture. Examination intervals vary and are dependent on target species, type of trap, weather, season, terrain, and number and experience of investigators. Generally, live traps for nocturnal species are set before dusk and checked at dawn. Traps are then retrieved or closed during the day to prevent capture of diurnal, nontarget taxa. However, live traps for small mammals, particularly shrews, should be checked more frequently (e.g., every 1.5 h—Hawes 1977) to minimize mortality due to the higher metabolism of these animals. Similarly, species of larger size with high metabolic rates (e.g., Mustela) also require shorter intervals between checking traps. Live traps for diurnal species should be set in areas shaded at dawn or early morning or under trap shelters (Kaufman and Kaufman 1989) and checked every few hours in warm weather.

 

Trap set
(Trap set in shade of shrub. This protects the trap from direct sunlight and overheating. Photo by Adam Green)

 

Traps should then be retrieved or closed at dusk to prevent accidental capture of nocturnal taxa. Thermoregulatory demands, especially for small mammals, can stress an animal even if duration of captivity is short. Thermoregulatory stress can be minimized by providing an adequate supply of food and nesting material in the live trap. Because most live traps for small mammals are constructed of metal and conduct heat readily, it might be necessary to insulate traps to minimize hypo- and hyperthermia in captive animals. Insulation can be accomplished by using such items as cotton or synthetic fiber batting, leaves, or twigs to provide dead air space between the animal and conducting surface and to provide escape from the temperature extremes. Critical temperature tolerance limits vary with species and environmental conditions. Investigators must be responsive to changing conditions and modify trapping procedures as necessary to minimize thermal stress.

If disturbance (removal of animal or trap damage) of live traps for small mammals by larger species of carnivores, birds, and others is problematic, trap enclosures (Getz and Batzli 1974; Layne 1987) or other methods to secure traps might be required.

Pitfall traps can be fitted with raised covers to minimize capture of nontarget species, provide cover from rain and sun, and prevent predation from larger animals. Pitfall traps used for live capture might require small holes in the bottoms to allow drainage in rainy weather, or enhancements such as small sections of polyvinyl chloride (PVC) pipe to provide escape from other captured animals.

Traps used for live capture of larger mammals include box traps, clover traps, and culvert traps. Some large mammals (e.g., ungulates and kangaroos) can be herded along fences into corrals or captured with cannon nets or drop nets projected from helicopters using net guns. These methods require immediate attention to the animals by trained personnel to prevent injury and can cause substantial distress in some species. With a large-scale capture, it may be useful to have a veterinarian on hand to assist with any injured or stressed animal. Depending on the nature of the work, individuals captured using these techniques may need to have their eyes covered or be sedated until the investigator’s work is completed (Braun 2005). Large mammals also can be captured by shooting a sedative into the hip or shoulder musculature using a dart gun. Baits laced with tranquilizer have been described (Braun 2005), but these should be used with caution to prevent sedating nontarget species. Chemical immobilization, whether for capture or sedation, requires training by a wildlife veterinarian and thorough knowledge of proper dosage, antidote, and sedative effect before use.

Judgment about use of local anesthetics when sampling peripheral body tissue and tissue fluids, such as blood, lymph, sperm, and tissue samples from body openings, should be based on a conscious effort to avoid or minimize pain and distress to the animal. If pain is slight or momentary, anesthesia is not recommended so that the animal can be released immediately. If pain is more than slight or momentary, field-portable anesthetic machines allow use of isoflorane and similar inhalants to provide a reliable anesthetic and rapid recovery after the animal is no longer exposed to the gas. Use of anesthesia for blood sampling will depend on data needed and species requirements. For example, some anesthetics (e.g., ketamine) depress blood pressure and make blood collection lengthier and potentially dangerous. Anesthesia also might alter the blood component (e.g., cortisols) under investigation. Use of anesthesia should be weighed against risk of mortality because some species are very sensitive to anesthesia (e.g., felids—Bush 1995; Kreeger 1996).

Investigators should bear in mind that stress and restraint associated with immobilization might be greater than applying or reading a particular mark without immobilization. Whether or not immobilization is required must be considered on a case-by-case basis. Procedures with animals that may cause more than momentary or slight pain or distress should be performed with appropriate sedation, analgesia, or anesthesia (article V, United States government principles for the utilization and care of vertebrate animals used in testing, research, and training; http://grants.nih.gov/grants/olaw/references/phspol.htm). Selection of anesthetics and analgesics for specific mammals should be based on evaluation by a specialist, such as a wildlife veterinarian, knowledgeable about the use of anesthesia in species of mammals other than standard laboratory or pet taxa. The investigator should conduct a literature review for alternatives as well as anesthetics and analgesics used in related species (Kreeger 1996). Physiological measurements required for experimental purposes also may affect the choice of anesthesia. Sedatives, anxiolytics, and neuromuscular blocking agents are not analgesic or anesthetic and hence do not relieve pain; these substances must be used in combination with a suitable anesthetic or analgesic (NRC 1996).

 

 

Measuring mouse
(Anaesthetized mouse immobilized for taking body measurements. Photo by Adam Green)

 

sedating mouse
(Sedating a mouse for measurements, collection of ectoparasites, and tissue sample for DNA analysis. Test tube with sedative can also be placed into a ziploc baggy with mouse. This should only be done by someone well trained in the tehnique because there is a fine balance between too little that results in a stressed and struggling animal and too much that can result in death. Photo by Adam Green)

 

Woodrat on back
(Woodrat aneasthetized for study on activity of detoxifying enzymes in the field. All woodrats recovered fully and were released back to their territories. Photo by Adam Green.)

 

An excellent reference for chemical immobilization of mammals is Kreeger (1996). Local and national regulations may restrict use of certain drugs (e.g., narcotics). Location of the animal within the habitat should be considered in light of time necessary for sedation and recovery to avoid injury or drowning of the sedated mammal. Further, sedated mammals must be monitored closely and observed after release until they regain normal locomotion. In no instance should sedated animals be left in proximity to water or exposed to potential predators while under the influence of immobilizing drugs.

Bats can be captured effectively and humanely with mist nets, harp traps, bag traps, or by hand (Kunz and Kurta 1988). Mist nets should not be left unattended for more than 15 min. Captured bats should be removed from nets immediately to minimize injury, drowning, strangulation, or stress. Removal of bats from mist nets must be done carefully to minimize stress and avoid injury to delicate wing bones and patagia. If a bat is badly tangled, it can be removed by cutting strands of the net. Mist nets should not be used where large numbers of bats might be captured at once, such as at cave entrances, because numbers can quickly overwhelm the ability of investigators to remove individuals efficiently. In these situations, harp traps or sweep nets are preferred (Wilson et al. 1996). Although harp traps do not require constant attention, they should be checked regularly, especially when a large number of captures is expected in a short period of time. Investigators using harp traps should guard against predators entering the trap bag, biting, predation of 1 bat species on another, rabies transfer, or suffocation due to large numbers of bats caught in a short time (Kunz and Kurta 1988).

As with traps for terrestrial mammals, to minimize stress on captured bats, the number of mist nets operated at one time should not exceed the ability of the investigators to check and clear nets of bats. Nets should not be operated in high winds because these conditions can put undue stress on bats entangled in nets. Mist nets should be operated only at night or during crepuscular periods and closed during the daytime to prevent capture of nontarget taxa (e.g., birds).

Roosting bats sometimes can be removed by hand. Gloves should be used that offer protection from bites but still allow the investigator to feel the body and movements of the bat to prevent injury to the animal. Long, padded tissue forceps might be used to extract bats from crevices, but extreme care should be taken to avoid injury to delicate wing bones and membranes (Kunz and Kurta 1988). The time of year that bats are studied must be considered and may be crucial to their survival. Large or repeated disturbance of maternity colonies might cause mortality of offspring and colony abandonment (Kunz 2003). Also, repeated arousal of hibernating bats can lead to mortality because of depletion of critical fat stores (Thomas 1995).

 

Woodrat restrained in plastic bag

 

 

 

Captured small and medium-sized mammals should be handled by methods that control body movements without restricting breathing.

 

 

 

(Woodrat restrained in plastic bag after capture. The handler has the woodrat restrained behind the neck. This immobilizes the rat and prevents it from biting the handler. It's important to quickly move the animal to another holding container or complete measurements and release. As can be seen here the rat is wet from perspiration in the plastic bag. Photo by Adam Green)

Covering an individual’s eyes might reduce the animal’s struggle to escape. Restraint by a mesh or cloth bag allows the investigator to mark, measure, or otherwise sample an individual through mesh or the partially opened end of the bag (e.g., Cynomys gunnisoni—Davidson et al. 1999).

 

woodrat in net funnel

(Use of a mesh cone to wpork with a woodrat. The mesh cone restrains the rat and allows a researcher to administer an injection. Photo by Adam Green)

Some small mammals also can be transferred directly from a trap to a heavy-duty plastic or cloth bag for transport.

 

mouse in bag
(By palcing a plastic bag over one end of the trap you can open the door of the trap and flip the animal into the bag without having to handle it. Some measurements and observations can be made while the animal is in the bag and the researcher can restrict the animal's movement to get a proper restraining hold for additional measurements. Photo by Adam Green.)

 

Design of some traps (e.g., box-type traps such as Sherman or Tomahawk live traps) also allows them to be used as a temporary cage for easy and safe transportation.

 

MARKING FOR IDENTIFICATION

Individual identification of mammals is necessary for many types of studies, both in the laboratory and field. Identification marks can be natural (stripe pattern, color, or mane patterns) or those applied by the investigator. Further, marks may be temporary or permanent, and external or internal. Of primary concern is the distance from which the animal must be identified. On large species, cataloging natural variations in fur or whisker patterns (West and Packer 2002), or previously sustained injuries on body parts (such as to wing, ears, or flukes) might be sufficient for permanent identification at a distance.

External Marks and Tags

Where naturally occurring identifying marks are not available, external dye, freeze-branding, or paint marks might provide the degree of longevity required. Dye marks on juveniles or subadults are of more limited duration because of rapid molting. Identification marks might be made with nontoxic hair dyes or paint. Care should be taken to ensure that substances used for external marks are nontoxic and do not otherwise alter the behavior of animals or subject them to increased predation. Freeze branding is an effective means of marking bats and other species and marks might last several years (Sherwin et al. 2002).

Metallic or plastic tags and bands or collars are cost-effective and might be suitable for identification at appreciable distance on large terrestrial species. Tags typically are applied to the ears of terrestrial mammals and to flippers of seals and sea lions. Use of individually numbered tags on small mammals necessitates handling the animal each time an individual is to be identified.

Although they are frequently used with a high degree of success, ear tags might inhibit grooming of ears and promote infection by parasites in some rodents (Ostfeld et al. 1996), although potential for infection likely varies with species and environment. Further, unless carefully sized, tags might snag, either during grooming or by vegetation in freeranging animals, and can be lost (Wood and Slade 1990). Many of the problems associated with ear tags are reduced in laboratory settings, where ear tags might be especially useful for long-term identification. Ear tags are not an option for species with greatly reduced pinnae (e.g., shrews).

Wing bands for bats should be applied so that they slide freely along the forearm, which may necessitate cutting a slit in the wing membrane in some cases. Another external marking option for bats is a bead-chain necklace (Barclay and Bell 1988), although these necklaces must be sized carefully.

 

ear punch
(Ear punch to collect tissue for DNA analysis. Photo by Adam Green)

 

 

Tattooing and ear punches provide a permanent means of identification but require handling of individuals for individual recognition.

Individuals of some taxa might be identified by unique patterns of ear punches (where a small amount of tissue is removed from external pinnae using some type of hole punch) or toe clips. Toe clipping involves removal of 1 or more digits (generally only 1 per foot) and provides a permanent identifying mark. Because both of these methods involve removal of a small amount of tissue, they might be especially appropriate in studies where tissues samples also are required.

Neither of these methods is generally suitable for identification at a distance, and ear punches might become unidentifiable through time in freeranging individuals because of healing, subsequent injuries sustained in the field, or being obscured by hair.

Because it is more invasive and addressed specifically in the Guide (NRC 1996), toe clipping requires considerable justification to the IACUC. Justification for toe clipping as a means of identification should include consideration of the natural history of the species, how the feet are used in the animal’s environment, and the size of the toe of the organism. Digits generally should not be removed from the forefeet of subterranean or fossorial taxa where they are used for digging. Nor should primary digits be removed from arboreal or scansorial taxa where they are used for climbing. Further, the size of the toe is related to body size of the animal; toe clipping in species with fleshy digits should be avoided. Toe clipping might be especially suitable for permanent identification in small species (e.g., Chaetodipus, Perognathus, Reithrodontomys, and Sorex) and in neonates of larger taxa. Toe-clipping and ear punches should not be used for marking bats; bats can be effectively wing punched or freeze branded. Toe clips and ear punches should be performed with sharp, sterilized instruments. Anesthetics and analgesics generally are not recommended; prolonged restraint of small mammals to apply these substances and consumption of the analgesic substances (e.g., creams) via licking likely cause more stress and harm than conducting the procedure without their use.

Radiotransmitters provide a mechanism to monitor movements and survival of individuals and, therefore, also serve to identify an individual. Transmitters can be attached externally with surgical or skin glue or a collar, or implanted into the animal’s body cavity. External attachment often can be accomplished in the field (e.g., Munro et al. 2006; Rothmeyer et al. 2002), whereas more invasive implantation might require transport to a laboratory where sterile conditions can be arranged. Investigators using collars should take into account potential for growth of an animal or seasonal changes in neck circumference (e.g., male cervids) and use devices designed to accommodate such changes (Strathearn et al. 1984). If external transmitters are attached using glue, individuals of some species will groom each other excessively to remove adhesive from their fur (Wilkinson and Bradbury 1988). Surgical implantation and more invasive procedures should be performed by a veterinarian or individuals who have received specialized training and usually require a suitable recovery period before the animal may be released.

Before using radiotransmitters, an investigator should consider the weight of the transmitter relative to the body mass of the target species or individual. Generally, the transmitter should represent, 5–10% of the individual’s body weight (Wilson et al. 1996).

As an alternative to radiotransmitters, light-emitting diodes (LED) or similar markers might be fastened externally to some species using similar considerations.

Internal Tags

Passive integrated transponder (PIT) tags are electronic devices encased in glass or resin capsules. They do not emit constant signals but can be interpreted with a remote reader in much the same way that bar codes are scanned. Tags are injected subcutaneously by using a modified large-bore hypodermic syringe and are suitable for many field and laboratory identification needs. Tags should be massaged away from the point of insertion subdermally to prevent loss. Even the smallest PIT tags (about the size of a grain of rice) may be too large for some individuals, so their use in very small individuals should be approached cautiously. Currently available PIT tag readers must be in reasonably close proximity to the tag (10 cm) for reading, so their use with large aggressive taxa (e.g., Procyon and Lynx) might require anesthesia both for application of the tag and for subsequent reading to prevent injury to the animal and investigators. Because of stress for both subject and investigator, other methods of tagging large mammals, such as using radiotransmitters or naturally occurring markers, might be preferable. Ingestion of colored plastic particles or radioactive isotopes (such as P32) in bait can be used to mark feces for studies of movements of individuals or groups of individuals, but is of limited utility to mark uniquely a large number of individuals.

 

HUMAN SAFETY

Working with wild mammals, particularly in field situations, comes with inherent risks, both biotic (e.g., bites, pathogens, parasites, and venomous plants and animals) and abiotic (e.g., lightning and exposure). Fortunately, most of these risks can be minimized with basic training, planning, mentoring, and experience. Investigators have the responsibility to ensure that personnel handling, transporting, or maintaining wild-caught mammals are qualified and familiar with requirements of the target species (e.g., bats - Constantine 1988) and hazards (e.g., bites and exposure to body fluids) associated with them. With appropriate preparation and training, investigators can adequately protect themselves and collaborators while conducting fieldwork with mammals (Kunz et al. 1997). Many universities and other institutions offer field courses, workshops, and online programs for investigators and students to achieve the proper training in fieldwork and in working with wild-caught mammals. Occupational health medical staffs also are available and aware of ways to avoid biological, chemical, and other hazards. Sources such as the Centers for Disease Control and Prevention (CDC 1998, 1999; http://www.cdc. gov/) or state health departments can provide current information for precautions against specific risks.

Special precautions to ensure human safety might be necessary when transporting taxa known or suspected of carrying potentially lethal pathogens such as Hantavirus or rabies. In areas where zoonotics are known to occur, bagging traps with a gloved hand and bringing them back to a central processing area that follows institutional biosafety recommendations might be sufficient. Additional precautions might be required at the time of final processing of the captured animal depending on data required. Although chloroform is considered highly hazardous to personnel with attendant health risks of cancer and liver toxicity (http://www.osha.gov/SLTC/healthguidelines/ chloroform/recognition.html), under open-air field conditions, its use might be appropriate because it kills ectoparasites that might pose greater risks to the researcher through transmission of diseases such as plague, typhus, or agents such as Hantavirus.

Many IACUCs will require the investigator to document their protocols for human health and safety while working with mammals. However, investigators and IACUC members should remain cognizant of the fact that risks from zoonoses vary depending on local environmental conditions, species of mammals handled, and the potential pathogens. Safety precautions should match perceived risks.

 

LITERATURE CITED

AD HOC COMMITTEE FOR ANIMAL CARE GUIDELINES. 1985. Guidelines for use of animals in research. Journal of Mammalogy 66:834.

AD HOC COMMITTEE ON ACCEPTABLE FIELD METHODS IN MAMMALOGY. 1987. Acceptable field methods in mammalogy: preliminary guidelines approved by the American Society of Mammalogists. Journal of Mammalogy 68 (Suppl.):1–18.

AMERICAN VETERINARY MEDICAL ASSOCIATION. 2001. 2000 report of the AVMA Panel on Euthanasia. Journal of the American Veterinary Medical Association 218:669–696.

ANIMAL CARE AND USE COMMITTEE. 1998. Guidelines for the capture, handling, and care of mammals as approved by the American Society of Mammalogists. Journal of Mammalogy 79:1416–1431.

ASPER, E. D. 1975. Techniques of live capture of smaller Cetacea. Journal of the Fisheries Research Board of Canada 32: 1191–1196

BAKER, R. J., AND S. L. WILLIAMS. 1972. A live trap for pocket gophers. Journal of Wildlife Management 36:1320–1322.

BARCLAY, R. M. R., AND G. P. BELL. 1988. Marking and observational techniques. Pp. 59–76 in Ecological and behavioral methods for the study of bats (T. H. Kunz, ed.). Smithsonian Institution Press, Washington, D.C. BOICE, R. 1972. Water addiction in captive desert rodents. Journal of Mammalogy 53:395–398.

BRAUN, C. E. 2005. Techniques for wildlife investigations and management. The Wildlife Society, Bethesda, Maryland. BUSH, M. 1995. Methods of capture, handling, and anesthesia. Pp. 25–40 in Wild mammals in captivity: principles and techniques (D. G. Kleiman, M. E. Allen, K. V. Thompson, and S. Lumpkin, eds.). University of Chicago Press, Chicago, Illinois.

CANADIAN COUNCIL ON ANIMAL CARE. 1993. Guide to the care and use of experimental animals (E. D. Olfert, B. M. Cross, and A. A. McWilliams, eds.). 2nd ed. Canadian Council on Animal Care, Ottawa, Ontario, Canada. Vol. 1.

CENTERS FOR DISEASE CONTROL AND PREVENTION. 1998. Hantavirus pulmonary syndrome—Colorado and New Mexico. Morbidity and Mortality Weekly Report 47:449–452.

CENTERS FOR DISEASE CONTROL AND PREVENTION. 1999. Update: hantavirus pulmonary syndrome—United States. Morbidity and Mortality Weekly Report 48:521–525.

CHOATE, J. R., AND H. H. GENOWAYS. 1975. Federal and state regulations pertaining to systematic collections. I. A case of inadvertent violation of federal regulations. SWANEWS (3&4): 10–13.

CONSTANTINE, D. G. 1988. Health precautions for bat researchers. Pp. 491–526 in Ecological and behavioral methods for the study of bats (T. H. Kunz, ed.). Smithsonian Institution Press, Washington, D.C.

DAVIDSON, A. D., R. R. PARMENTER, AND J. R. GOSZ. 1999. Responses of small mammals and vegetation to a reintroduction of Gunnison’s prairie dogs. Journal of Mammalogy 80:1311–1324.

DIERAUF, L. A., AND F. M. D. GULLAND (EDS.). 2001. CRC handbook of marine mammal medicine. 2nd ed. CRC Press, Boca Raton, Florida.

FOWLER, M. E. 1995. Restraint and handling of wild and domestic animals. Iowa State Press, Ames. GETZ, L. L., AND G. O. BATZLI. 1974. A device for preventing disturbance of small mammal live-traps. Journal of Mammalogy 55:447–448.

GREER, L. L., J. WHALEY, AND T. K. ROWLES. 2001. Euthanasia. Pp. 729–740 in CRC handbook of marine mammal medicine (L. A. Dierauf and F. M. D. Gulland, eds.). 2nd ed. CRC Press, Boca Raton, Florida.

GRINNELL, J. 1928. Recommendations concerning the treatment of large mammals in Yosemite National Park. Journal of Mammalogy 9:76.

HAFNER, M. S., W. L. GANNON, J. SALAZAR-BRAVO, AND S. T. ALVAREZCASTAN ˜ EDA. 1997. Mammal collections in the Western Hemisphere: a survey and directory of existing collections. American Society of Mammalogists, Allen Press, Lawrence, Kansas.

HART, E. B. 1973. A simple and effective live trap for pocket gophers. American Midland Naturalist 89:200–202.

HAWES, M. L. 1977. Home range, territoriality, and ecological separation in sympatric shrews, Sorex vagrans and Sorex obscurus. Journal of Mammalogy 58:354–367.

KAUFMAN, D. W., AND G. A. KAUFMAN. 1989. Burrow distribution of the thirteen-lined ground squirrel in grazed mixed-grass prairie: effect of artificial habitat structure. Prairie Naturalist 21:81–83.

KREEGER, T. J. 1996. Handbook of chemical immobilization. International Wildlife Veterinary Services, Inc., Laramie, Wyoming.

KUEHN, D. W., T. K. FULLER, L. D. MECH, W. J. PAUL, S. H. FRITTS, AND W. E. BERG. 1986. Trap-related injuries to gray wolves in Minnesota. Journal of Wildlife Management 50:90–91.

KUNZ, T. H. 2003. Censusing bats: challenges, solutions, and sampling biases. Pp. 9–17 in Monitoring trends in bat populations of the United States and territories: problems and prospects (T. J. O’Shea and M. A. Bogan, eds.). United States Geological Survey Information and Technology Report, USGS/BRD/ITR-2003-003. National Technical Information Service, Springfield, Virginia.

KUNZ, T. H., AND A. KURTA. 1988. Capture methods and holding devices. Pp. 1–29 in Ecological and behavioral methods for the study of bats (T. H. Kunz, ed.). Smithsonian Institution Press, Washington, D.C.

KUNZ, T. H., R. RUDRAN, AND G. GURRI-GLASS. 1997. Human health concerns. Appendix 2. Pp. 255–264 in Measuring and monitoring biological diversity: standard methods for mammals (D. E. Wilson, F. R. Cole, J. D. Nichols, R. Rudran, and M. S. Foster, eds.). Smithsonian Institution Press, Washington, D.C.

LAYNE, J. N. 1987. An enclosure for protecting small mammal traps from disturbance. Journal of Mammalogy 68:666–668.

LEACH, M., V. BOWELL, T. ALLAN, AND D. B. MORTON. 2002. The aversion to various concentrations of different inhalational general anesthetics in rats and mice. Veterinary Record 150: 808–815.

MACRINA, F. L. ( ED.). 2005. Scientific integrity: text and cases in responsible conduct of research. 3rd ed. American Society of Microbiology Press, Washington, D.C.

MARTIN, R. E., A. F. DEBLASE, AND R. H. PINE. 2000. A manual of mammalogy: with keys to families of the world. McGraw-Hill, New York.

MUNRO, R. H. M., S. E. NIELSEN, M. H. PRICE, G. B. STENHOUSE, AND M. S. BOYCE. 2006. Seasonal and diel patterns of grizzly bear diet and activity in west-central Alberta. Journal of Mammalogy 87:1112–1121.

NATIONAL RESEARCH COUNCIL. 1992. Recognition and alleviation of pain and distress in laboratory animals. National Academy Press, Washington, D.C.

NATIONAL RESEARCH COUNCIL. 1996. Guide for the care and use of laboratory animals. National Academy Press, Washington, D.C.

NATIONAL RESEARCH COUNCIL. 2003. Guidelines for the care and use of mammals in neuroscience and behavioral research. National Academy of Sciences, Washington, D.C.

OFFICE OF LABORATORY ANIMAL WELFARE. 2002. Institutional animal care and use committee guidebook. 2nd ed. National Institutes of Health, Department of Health and Human Services, Bethesda, Maryland.

OFFICE of LABORATORY ANIMAL WELFARE/NATIONAL INSTITUTES of HEALTH. 2002. Public Health Service policy on human care and use of laboratory animals. National Institutes of Health, Bethesda, Maryland.

OSTFELD, R. S., M. C. MILLER, AND K. R. HAZLER. 1996. Causes and consequences of tick (Ixodes scapularis) burdens on whitefooted mice (Peromyscus leucopus). Journal of Mammalogy 77: 266–273.

POWELL, R. A., AND G. PROULX. 2003. Trapping and marking terrestrial mammals for research: integrating ethics, performance criteria, techniques, and common sense. Institute for Laboratory Animal Research Journal 44:259–276.

ROTHMEYER, S. W., M. C. MCKINSTRY, AND S. H. ANDERSON. 2002. Tail attachment of modified ear-tag radio transmitters on beavers. Wildlife Society Bulletin 30:425–429.

RUSAK, B., AND I. ZUCKER. 1975. Biological rhythms and animal behavior. Annual Review of Psychology 26:137–171.

SCHMINTZ, S. D. 2005. Capturing and handling wild animals. Pp. 239–285 in Techniques for wildlife investigations and management (C. E. Braun, ed.). The Wildlife Society, Bethesda, Maryland.

SHERWIN, R. E., S. HAYMOND, D. STRICKLAN, AND R. OLSON. 2002. Freeze branding for permanently marking temperate bat species. Wildlife Society Bulletin 30:97–100.

SOCIETY FOR THE STUDY OF ANIMAL BEHAVIOUR. 2006. Guidelines for the treatment of animals in behavioural research and teaching. Animal Behaviour 71:245–253.

STRATHEARN, S. M., J. S. LOTIMER, G. B. KOLENOSKY, AND W. M. LINTACK. 1984. An expanding break-away radio collar for black bear. Journal of Wildlife Management 48:939–942.

THOMAS, D. W. 1995. Hibernating bats are sensitive to nontactile human disturbance. Journal of Mammalogy 76:940–946.

UNITED STATES DEPARTMENT OF AGRICULTURE. 2005. Animal Welfare Act and animal welfare regulations. Animal Welfare Act as of November 1, 2005 as found in the United States Code, Title 7— Agriculture, Chapter 54—Transportation, Sale, and Handling of Certain Animals, Sections 2131–2159. United States Government Printing Office, Washington, D. C.

WALKER, W. A. 1975. Review of the live-capture fishery for smaller cetaceans taken in southern California waters for public display. Journal of the Fisheries Research Board of Canada 32:1197–1211.

WEST, P. M., AND C. PACKER. 2002. Sexual selection, temperature, and the lion’s mane. Science 297:1339–1343.

WILKINSON, G. S., AND J. W. BRADBURY. 1988. Radio telemetry: techniques and analysis. Pp. 105–124 in Ecological and behavioral methods for the study of bats (T. H. Kunz, ed.). Smithsonian Institution Press, Washington, D.C.

WILSON, D. E., F. R. COLE, J. D. NICHOLS, R. RUDRAN, AND M. S. FOSTER (EDS.). 1996. Measuring and monitoring biological diversity: standard methods for mammals. Smithsonian Institution Press, Washington, D.C.

WILSON, D. E., AND D. M. REEDER (EDS.). 2005. Mammal species of the world: a taxonomic and geographic reference. 3rd ed. Johns Hopkins University Press, Baltimore, Maryland. Vols. 1 and 2.

WOOD, M. D., AND N. A. SLADE. 1990. Comparison of ear-tagging and toe-clipping in prairie voles, Microtus ochrogaster. Journal of Mammalogy 71:252–255.

 

Species caught at Coal Oil Point Feb 6, 2015:

mus vs rhythrodonimes
(Left: Mus ; Right Reithrodontomys. Photo by Adam Green)
MicrotusPeromyscus
(Left: Microtus; Right: Peromyscus. Photo by Adam Green)

Some good resources for identifying mammals:

Discover Life_Mammalia


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