The overall goal of this procedure is to utilize a natural vector, the deer tick for experimental transmission and xeno diagnosis of the Lyme disease agent Borrelia Borre. This is accomplished by first producing ticks that harbor the bacteria. Once inoculated, the ticks are applied to the shaved area of a mouse and then housed in specialized caging.
Fed ticks are then retrieved from the cage water. For non-human primates, a specialized containment device is constructed and applied before the ticks are placed on the skin. After five days, the devices opened and fed ticks are collected.
Ultimately, samples can be collected and analyzed by various methods to confirm infection after tick. Feeding tissues can also be cultured, sectioned, and stained for immunofluorescence. Visual demonstration of these methods is critical as the multiple steps involved are actually quite different between the animal models used.
Demonstrating the procedure will be myself, Dr.Britton, grass Bears, and Mary Jacobs White lab coats with elastic sleeves, gloves, and disposable bouffant caps should be worn when working with ticks. To begin retrieve nimal ticks that have been stored at 23 degrees Celsius for four to six weeks. Post larva molt place, double-sided tape on the bottom of a small Petri dish and secure the ticks ventral side up on the tape for inoculation.
Beberg Dory are grown to midlock phase in BSKH medium containing 6%rabbit serum glass capillary tubes are prepared using a tube standardized to the tick mouth part as a sizing guide. When ready, mix the Beberg Doy culture and insert the tip of the capillary tube With the aid of a dissecting scope, place the tube over the hyperdome of the tick mouth. Next, fix the tube in place with molding clay.
Insert the Petri dish with a fixed ticks inside a large clear plastic tub for an added level of containment and add wet paper towels to provide moisture. Place the ticks in a 37 degree Celsius incubator until defecation is apparent. This indicates that the media containing spirochetes has passed through the tick.
All the ticks are kept for two to four weeks at 23 degrees Celsius to adapt to their environment before feeding them on animals. After administering anesthesia, shave the mouse from the ears to the middle back, using an electric trimmer in a white pen with no other objects nearby. Use a moistened paintbrush to transfer the inoculated ticks to the hairless area of the mouse.
Once the ticks are in place, transfer the animal to a specialized cage. This cage consists of a stainless steel grill elevated from the bottom. The cage top was modified in-house to elevate the water bottle holder enough to allow free movement of the mouse underneath to trap any ticks that may fall from the mouse.
The cage bottom is filled with just enough water to cover the surface, but is low enough to prevent limbs from submersion. Heating pads are placed underneath the cage until the mouse is fully recovered. The cage is also placed inside a tray that has been lined with tangle tre paste and tape for further measures of containment.
When the mouse has fully recovered from anesthesia, water is poured into the cage to a depth of about one half inch, and food and water are replaced after three, four, and five days. Check the mouse cage and cage water for fed ticks. Sift the cage water through a white metal pan.
Rinse the fed ticks in clean water and store them in plastic jars on days three and four. Replace the water in the cage with clean water. On day five, check not only the cage, but the mouse thoroughly for ticks.
Usually by this point, all ticks have fed and the mice can be returned to regular caging. Place all waste from the mouse cages, including liquids in biohazard containers. For autoclaving and disposal, keep a log of the number of ticks placed on the mice and those recovered at all times.
The first step is to prepare the tick containment device. Cut a one and three quarter inch diameter circle in the three by three inch LA flap using a clean scalpel and the measurement guide, use the cutout as a template to cut circles of identical size in the biota foam and DuoDERM. The foam is used to elevate the flap over the surface of the skin and prevent possible crushing of the tick.
The duo derm adds another layer of cushioning and overlays the edges of the containment device for added security from tick escape. After administering anesthesia, an electric trimmer is used to shave all the areas that will be covered by the jacket. Next, closely shave an area of approximately 25 by 20 centimeters.
Then clean and dry the area. After cleaning and drying the skin place the duo derm on the animal's dorsum just below the scapula on either side of the spine. Use a marker to trace the circle in that spot.
Skin prep is used to wipe skin to remove oils. After wiping the skin, apply a layer of skin glue. Remove the adhesive backing from the biotine foam and affix it to the skin in the appropriate spot.
Seal the edges with skin glue and type fixx tape. Remove the adhesive backing from the la flap and fix on top of the Biotine Place skin glue and hyper fixx Tape around the edges. After taping down the mesh flap, place the jacket on the animal.
24 hours later, check the security of the device and reinforce if needed. When the device is secure, use a paintbrush to place approximately 20 unfed ticks on the skin. After transferring the ticks, remove the adhesive backing from the mesh of the flap and seal it in place.
Remove the duo derm backing and place it on top of the containment device. Seal the edges with hyper fixx tape. Lastly, add a piece of hyper fixx tape across the open mesh circle and replace the jacket.
Five days later, the animal is anesthetized and the jacket is removed. Remove the tape first so that tick feeding can be viewed through the mesh. After carefully peeling the duo derm away from the flap, pull the mesh portion back at the edges to provide access to the ticks.
Fed ticks are frequently found near or under the foam circle. Remove the ticks with a paintbrush and place them in clean water. Once all visible fed ticks are collected, remove the device and wipe the skin with alcohol.
Finally, store the ticks at 23 degrees Celsius for future use. Using the capillary feeding technique, over 90%of Fed ticks are found to harbor bedoy as demonstrated here by antibody staining and fluorescent microscopy. Mouse infection rates with a low passage strain are near 100%A combination of serology as shown here and culture are used to determine if each mouse has become infected While attempting the capillary feeding procedure.
It's important to remember to keep checking the ticks periodically for evidence of feeding or displacement of tubes from the mouth parts After their development. The techniques for feeding ticks on mammals paved the way for researchers in the field of wine disease to explore both host and pathogen responses following natural infection. After watching this video, you should have a good understanding of how to generate ticks of the ex exotic species that harbor bor and fe ticks on Meer monkeys for studies of Lyme disease.
