Flea Collection Techniques

For small rodents, e.g., Peromyscus, Neotoma, Apodemus, etc. small collapsible aluminum Sherman traps (3 x 3 x 10 inch) work well using oatmeal for bait. Baits such as seeds, fruits, tubers, etc, may be used in different localities to match the natural food source of the animals being trapped. For nocturnal animals, traps are picked up right after first light and for diurnal animals about 10:00-11:00 A.M. (to avoid heat problems). White cloth bags (approximately 9 x 12 inches) sewn from a close weave material work well to contain small animals. White enables one to see the fleas that WILL be in the bags. As trap lines are checked, place the bag directly over the end of the trap, while pushing the door opened, shake the rodent into the bag. This gets fleas that might be in the trap as well as those remaining on the animals. Tie the sack closed with a string and immediately sacrificed the rodent by several methods: 1) by compressing the heart/lung area between the thumb and fingers or 2) cervical fracture. These are quick, bloodless, and do not damage the skull or study skin for later museum preparation. The animals will quickly chew out of the bags if not sacrificed. Then proceed to the next trap. Once finished, process the animals in the field or in the lab (depending on locality). If you use the catch and release methodology, leave the animals in the traps while transporting to an area to process them for fleas. Animals may also be euthanized with halothane or metafluorane, however, these gases also knock down the fleas and they are difficult to remove from the fur unless they are thoroughly washed (not usually practical in field situations).

To process small mammals, the white bag and all contents are poured into a five-gallon bucket (clean and white). The bag should be held down in the bucket while turning it inside out. Examine the surface for fleas, which can be flicked off into the bucket. Pick the animal from the bottom of the bucket and run a gentle stream of compressed CO2 over the fur for about 10-15 seconds while holding down in the bucket. The CO2 passes through 1/2 rubber tubing to a tip (1/2 inch copper pipe flared our by hammering one end relatively flat). The flanged tip delivers a broadly distributed stream of CO2 as you run it over the fur (as using a vacuum...except blowing a stream of CO2). Most fleas are agitated by the CO2 and vacate while in the white bucket. Some are less sensitive to CO2 and must be "combed" out. Combing is a practical alternative to using CO2. To comb the rodent, use an old toothbrush with all the rows (except for one or two rows) of bristles cut off flush with a razor blade. Pay particular attention to combing the neck, head and rump area over and around the tail. Frisk them briskly with the toothbrush. Fleas are highly adapted to hanging on and crawl deep in the fur. Some animals are unable to preen the nap of the neck and over the tail, so fleas often congregate in these areas.

Once satisfied that the animal no longer has any remaining fleas, place in a bag for mammal processing. Fleas may be examined immediately using a dissecting microscope or at a later date. For immediate examination, place a dish of alcohol (syracuse dish) in the bottom of the bucket and use an applicator stick (or any stick) sharpened to a fine point to pick up the fleas. Dip the stick into the alcohol and the evasive fleas will adhere to the stick. Transfer each flea to the syracuse dish. If you are not interested in examining the flea, use a vial instead of the syracuse dish. 70-80% ethanol adequately preserves fleas. Avoid using any concentrations of isopropyl or methyl alcohol, and higher concentrations of ethanol, since they will make fleas brittle. After fleas are examined with a dissecting microscope, place them in a vial (all the fleas from ONE animal in ONE vial). Place a duplicate field number in both the host animal bag and INSIDE the alcohol vial of fleas to associate the parasites with the voucher host specimen. Use a lead pencil or indelible ink to avoid accidental smudging or loss of data

The data associated with each field number should include (as an example):

Host animal species: Peromyscus yucatanensis
Sex of host animal: Male/Female
Developmental stage: Juvenile/Adult
Locality: 2 km NE of Merida, Yucatan, Mexico (include coordinates, or township, range, section data when possible, e.g., 45°30'15"N, 111°30'22"W, or T10S R9W Sec 33, etc.)
Date of collection: Day/Month/Year, e.g., 3 July 1998 (always write month out, OR use Roman numerals, e.g. VII; avoid using numbers for date, e.g., 3/7/98, or 7/3/98!)
Collector: J.Q. Doe

Habitat data, e.g., terrain features, vegetation types, etc., are valuable to associate with host-parasite relationships. Mammal measurements might also be recorded.

Ultimately, record data into a computerized format, e.g., Excel. This makes record keeping much easier and is essential for large collections in order to manipulate and manage the data. Remember: always leave a blank column for ectoparasite identifications. Other ectoparasites might also be collected and stored in the same vials for later sorting and distribution to specialists, i.e., ticks, mites, bat flies, etc.

Larger animals such as Armadillos, Conepatus, Didelphis, Vulpes, etc. must be examined visually over the entire body. CO2 will also work to flush the fleas from the hairy areas of these animals. This can be done while the animals are inside the cage as it is held over a white sheet. As the fleas jump off, they can be collected from the white sheet. Cages can also be placed in large plastic bags while CO2 from a cylinder or from a short blast from a CO2 fire extinguisher directed into the bag. Fleas may be recovered from the plastic bag. Animals should be anesthetized with an injection of ketamine hydrachloride before using CO2 in a closed space. Metaphane or halothane will euthanize not only the animals, but also the fleas, necessitating a post-treatment examination for dead fleas on the animals.