Collecting
For beginners, I would recommend trying lichens from rocks or trees first. These habitats will have less soil particles and the tardigrades will be easier for a neophyte to see and isolate. Moss, lichen, or soil habitat samples are best collected dry. When collected, place the habitat samples in small paper bags. Paper bags allow moisture to escape; plastic bags would trap moisture and possibly cause deterioration of the specimens. Plant material that is not dry can be placed in the paper bags, which should then be placed in an open, dry place to allow slow drying. Write collection data on the paper bag with pencil. Specimens can be obtained from samples such as this years later, although the older the sample the less probability that the tardigrades will be alive. In one location it is best to sample all of the species of likely habitat, as well as different exposures. Collecting tardigrades is a lot like photography -- the more samples taken, the better the chances of getting good results.
Extraction from Habitat Sample
If there are very few soil particles associated with the sample (e.g., lichens from a tree), a simple procedure can be used. Place plant material samples in a fingerbowl and add enough water to moisten all of the plant material. I use bottled spring water, but tap water will work (with less probability of the tardigrades staying alive). After 2-24 hours, swish the plant material around in the water and then remove the plant. (To identify some species eggs are needed, which may require the longer soaking time.) Examine the bottom of the fingerbowl with a dissecting microscope. At first, everyone has trouble finding tardigrades but it gets easier with experience. Remove the specimens with an Irwin loop or a micropipette.
Soil samples or moss samples with soil around the rhizoids are more difficult. It is possible to do as above and then slowly move around the soil particles with a dissecting needle and pick out the tardigrades. This consumes large amounts of time. Another approach is to use a floatation-centrifugation technique. I use a technique from Jenkins (1964), incorporating some of Roberto Bertolani's suggestions. I use a set of sieves, obtained from one of the larger scientific supply companies. The mesh can be changed in these. I use 180, 120, and 90 micron mesh sizes, which came with the set of sieves. In a fourth sieve, I place 45 micron mesh which was purchased separately and cut to fit the sieve. After soaking the sample, remove the larger solid material with a coarse sieve (mine is made with window screening). Place the sample in a centrifuge tube (I use 50 ml tubes). Centrifuge at 2,000 rpm for 5 minutes. Decant the liquid into the sieves. Add a sucrose solution (Place 70 ml of water in a graduated cylinder and add sucrose until total volume is 100 ml). Centrifuge at 3,000 rpm for one minute. Decant into the sieves. Rinse the sieves with water to prevent/reverse the shriveling of the tardigrades by the sucrose solution. Back-rinse the sieves into fingerbowls and examine with a dissecting microscope. Rarely would a tardigrade be found in the first sieve (180 micron mesh) but it removes the larger particulate matter. The tardigrades are found in the 120 and 90 micron mesh sieves, depending on the size of the tardigrade. Eggs will mainly show up in the last sieve. Using a polyethylene wash bottle, I backwash each sieve into a separate fingerbowl.
The
sieves can be used without the floatation-centrifugation technique. After soaking the sample, agitate the plant
material, then remove with forceps or a very coarse sieve. Then filter the liquid through the set of sieves, followed by backwashing
the sieves into separate fingerbowls for examination.
Mounting Media and Ringing Slides
References
Jenkins, W. R. 1964. A rapid centrifugal-flotation technique for separating
nematodes from soil. Plant Disease Reporter 48:692.
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