Directions for isolation
Isolating and Culturing Batrachochytrium dendrobatidis
Joyce E. Longcore, School of Biology & Ecology
University of Maine, Orono, Maine, 04469-5722 USA
Collection of Amphibians
Collect dead, moribund or suspect amphibians. If amphibians are killed or found dead, cool as soon as possible and keep cool and damp but not waterlogged until examination and possible isolation attempts. B. dendrobatidis can most easily be isolated from the mouth parts of tadpoles. Collect tadpoles with focal dekeratinized areas of the jaw sheaths, which can be detected in the field with a 10 X hand lens. Isolation of B. dendrobatidis should be attempted from skin samples from freshly dead frogs; the longer an animal is dead, the more problems will be encountered from bacteria and non-target fungi. For a non-lethal method to isolate from foot skin webbing see Fisher et al. [Scientific Reports volume 8, Article number: 7772 (2018)].
Recognition of Chytridiomycosis
First, microscopically identify B. dendrobatidis in fresh host tissue. Keep amphibian tissue damp, but without free water. While observing with dissecting microscope at 20 or 40 X, use a sharp, sterile needle to remove loose skin from between digits of foot and elsewhere on the ventral surface of the animal. If skin is not loose, use needle-nosed forceps, micro-scissors or a single-edged razor blade to remove pieces from the leading edge of the skin between the hind digits.
If isolating from larval animals, look for a focal lack of melanin of the jaw sheath (Fig. 1) and remove this area of the sheath with needle-nosed forceps. Place the skin or jaw sheath on a microscope slide in a drop of sterile distilled water and cover with a coverslip.
Observe with a compound microscope with 40 and 100 X objective lenses. Some sporangia may have cleaved zoospores, which will look like 5–20 spherical bodies within the sporangia. Other sporangia may have discharged their spores and appear empty. Look for walled, spherical to oval bodies (10–30 µm diam) inside of epidermal cells. Some of the bodies may be septate (delicate walls divide the fungal body, or thallus, into 2 or more sporangia).
If suspect organisms are found, place that piece of skin on a 9 cm culture plate containing mTGh or 1% tryptone nutrient agar plus antibiotics, which are added after autoclaving (see recipe below). While observing through a dissecting microscope (40 X magnification with substage lighting) use a sharpened and sterilized needle to draw and push a small (1 X 1 mm or less) piece of infected skin through the nutrient agar. (If pieces of skin are thick and do not tear into small pieces, use micro-scissors or cuticle scissors to cut skin into small pieces while viewing with 20 X magnification.) Every few millimeters take the needle away from the piece of skin and wipe through the agar. The purpose of wiping the skin and needle through agar is to remove bacteria, yeast and fungal spores. Bearing this in mind, reverse the direction of the skin, wipe it back and forth; imagine what you are trying to do even though you cannot see the bacteria. Jaw sheath tissue from recently killed larvae is usually nearly free of bacteria and other fungi and requires less cleaning. When the skin is well wiped (at least back and forth across the diameter of the plate one time), use the cleaned needle to remove the skin from the cleaning plate, which has been open to the air, and put it on a fresh plate of nutrient agar. Barely open the fresh plate with one hand and wipe the skin piece into the agar with the needle held in the other hand. This is so that the pieces of skin will be on a plate that has not been exposed to fungal spores from the air. Repeat this process for as many pieces of skin as you have the patience; at a minimum, for each isolation attempt, put 6 pieces of wiped skin on each of two plates. Seal the nutrient agar plate with a 10 X 2 cm piece of Parafilm® or other laboratory film stretched around the circumference of the plate. Label the dish with a permanent marker, recording source of skin and date; circle on the bottom of the plate areas where pieces of skin are located. Incubate sealed plates on the laboratory bench or, if laboratory temperatures are above 25o C, in an incubator at 17–23o C.
During the next week to month, check development by inverting the plate on the stage of a compound microscope and observing the small pieces of skin with the 10 X objective. Also, inspect plates for contaminants. If fungal contaminants are found, flame-sterilize a scalpel and remove the areas of contamination. Motile B. dendrobatidis zoospores (4–5 µm dia.) may be evident around the cleaned, infected skin within 1–2 days, but sometimes not for several weeks. If chytrid colonies develop on the skin, thalli can be recognized as spherical bodies, some of which bear one or more nipple-like, discharge papillae.
If hyphae are evident instead of round sporangia, use a sterile knife to aseptically remove the hyphal colony from the isolation plate. If B. dendrobatidis colonies are produced on many pieces of skin, one piece can be removed to examine the fungus with the compound microscope. Using techniques to minimize the chances of entry of airborne fungal spores, open a plate and to remove a piece of the colony with a sterilized scalpel and place in a drop of water on a microscope slide. Observe with the compound microscope and compare morphology with photographs of B. dendrobatidis, e.g., Fig 6.
After the chytrid forms a colony on the isolation plate, aseptically transfer a piece of the colony to a plate of 1% tryptone agar (without antibiotics). B. dendrobatidis grows best in groups of thalli, so do not separate sporangia from each other during transfer. Incubate for 1 or 2 weeks; if no bacteria develop and the fungus is growing well, spread the colony on the plate and transfer a part of the colony to nutrient broth in a screw-capped, 250 mL flask. For back-up, also transfer bits of the colony to fresh plates, seal with Parafilm®.
For stocks, keep 2 sets of cultures in 1% liquid tryptone; after growth is visible refrigerate for ~3 months between transfers. Stock cultures of most other chytrids are kept on agar slants in screw-capped test tubes; however, B. dendrobatidis survives longer in liquid medium. If stocks are kept on nutrient agar, refrigerate sealed Petri plates that contain scattered, small colonies of B. dendrobatidis.
Maintaining Cultures of B. dendrobatidis
Place 75 ml of 1 % liquid tryptone medium into 150 ml screw-capped flasks. The size of the flask or tube is not important – use larger or smaller containers depending on what is available. Because of the danger of contamination, use screw-capped vessels and keep duplicates. Incubate at 23o C or below. For long-term storage place at 5o C after growth is evident on the walls of the culture vessel. To observe growth macroscopically, slowly tip the container while watching the previously liquid-covered glass wall.
A reliable method to freeze and then store cultures in liquid nitrogen has been developed [Boyle, D. G. et al. (2003). Cryo-archiving of Batrachochytrium dendrobatidis and other chytridiomycetes. Diseases of Aquatic Organisms, 56, 59–64]. Cryopreservation allows isolates to be stored without the need to passage every 3 months, which is tedious and may result in contamination or changes in pathogenicity.
Production of Zoospores
Grow B. dendrobatidis in 1% liquid tryptone until clumps of thalli are visible to the eye. Use a sterilized Pasteur pipette to add 1/2 to 3/4 mL of this broth culture to 1% tryptone agar in 9 cm culture dishes. Leave inoculated dishes open in laminar flow hood until the added broth is dry; replace covers on dishes and place in plastic sleeve. Incubate in 15o C incubator. Plates can be incubated at higher temperatures, up to 23o C, but the potential harvest period is longer if plates are kept in plastic sleeves at 15o C. After 3–10 days look for active zoospores around the periphery of fungal colonies by inverting dishes on the stage of a compound microscope and examining with the 10 X objective.
Harvest zoospores by flooding plates with 2–3 mL of sterile, distilled water. Decant after about 30 minutes to collect zoospores. Zoospore concentration can be measured by optical density or by counting a measured amount on a hemocytometer. Zoospores may stay motile (thus infective) for up to 24 hours, however, most encyst before 24 hrs.
Although thorough drying can kill B. dendrobatidis, take all precautions. Before disposal, autoclave all materials that contain or have come into contact with the fungus. If B. dendrobatidis is used to inoculate amphibians, kill and incinerate or fix all exposed animals after the experiment. Be sure that cages, water, and other material in cages are disinfected at the end of experiments. Do not place potentially contaminated material in the trash or empty liquid in the drain without first treating it (heat or bleach) to kill the fungus.
Fig. 7. Colony of B. dendrobatidis surrounded by zoospores on plate of nutrient agar. Viewed by inverting on stage of compound microscope; 10 X objective lens.
mTGh Isolation medium
(add 200 mg/L penicillin-G and 200-500 mg/L
streptomycin sulfate after autoclaving; if bacteria are still a problem add 1 mg/L ciprofloxacillin)
8 g tryptone
2 g gelatin hydrolysate
10 g agar
1000 mL distilled water
1% Tryptone agar (for broth, delete agar)
10 g tryptone
10 g agar
1000 mL distilled water