Protocol for Ciliates and Testate Amoebae

Protocol for collecting and staining ciliates and testate amoebae from phytotelmata or edaphic samples

DIMARIS ACOSTA-MERCADO
University of Puerto Rico, Mayagüez Campus

  1. Use preliminary data to evaluate the spatial distribution of protists to avoid autocorrelated samples. This information is useful in designing a sampling strategy that captures a robust estimate of diversity with appropriate number of replicates (see Acosta-Mercado and Lynn, 2002).
  2. For aquatic samples, insert the pipette until reaching the bottom of the phytotelmata and gently stir the debris, collect 2.0 mL of that resuspended material. For edaphic ciliates and testate amoebae, place 20-30 g of freshly collected organic soil in a Petri dish. Remove large debris only (i.e. large >10 mm rocks, branches etc.). Saturate with distilled water until a runoff of 2.0 ml is obtained when the Petri dish is tilted at 45° (Foissner, 1987). Store Petri dishes at room temperature (~20-24ºC) and visually inspect to corroborate that moisture content is kept at a constant level. At Day 2 after saturation, move the Petri dish with a gentle rotary motion and then remove 0.5 ml from each of the four quadrants. Replace water removed with 2.0 ml of distilled water or filtered mineral water.
  3. Fix each sample with concentrated Bouin’s fluid (Coats and Heinbokel, 1982) to a final ratio of fixative to sample 1:20 (i.e. 100 :l of concentrated Bouin’s to 2.0 ml of run off). Note: Concentrated Bouin’s is made by saturating liquid 37% formaldehyde with calcium carbonate crystals, decanting, and then re-saturating the buffered formaldehyde with picric acid crystals. Before adding the runoff from the Petri dish, add glacial acetic acid to the Bouin’s fixative so that its final concentration, once the runoff is added, will be 1% (v/v) (i.e., 20 :l for 2.0 ml runoff). Let the samples remain in fixative at least for 48 h. Another alternative is to fix with 2-5% Acid Lugol’s fixative; however, these samples should be resuspended and postfixed with concentrated Bouin’s (5 % final concentration) before they can be processed (Montagnes & Lynn 1993).
  4. Gently invert the vials containing the fixed samples (Bouin’s or Lugol’s Fixative) (3-4 times) to release the cells from the soil substrates, and let these soil particles settle for 40 sec. Filter each sample through a SARTORIUS gridded cellulose nitrate BKL filter, 1.2 μm pore size (# 11403.025N) in a Millipore column. Apply suction of no more than 100 mm Hg to avoid rupturing cells (Montagnes and Lynn, 1993). Do not let the filters dry in the column. Wash them in the column several times with distilled water until the yellow Bouin’s color is reduced.
  5. Remove the filter and place it “sample side up” on a piece of warm glass.
  6. Place a drop of 2.5 % (w/v) agar in distilled water on a 20 X 30 mm coverslip, invert it, and quickly but gently place it on top of the filter on the glass. Apply slight pressure to assure that the agar forms a homogeneous and thin layer on top of the filter.
  7. Trim the two edges of the filter that extend past the coverslip, and peel off the filter. Place filters back to back in the first Columbia staining jar (A. H. Thomas, Inc., P.O. Box 779, Philadelphia, P. A., USA, 19105-0779), which has been filled with distilled water. Note: all solutions should be placed in the staining jars as the protocol starts.
  8. Bleach with 0.5% (w/v) of potassium permanganate (KMnO4) for 8 min. Ciliates from soil samples require more time at this stage presumably due to the presence of fumic acids, and other secondary compounds in the soil that reduce the efficiency of the permanganate.
  9. Wash the filters in running tap water for 5 min until the excess of KMnO4 is removed. Transfer the filters from one jar to another until the purple colour is removed.
  10. Place the filters in 5% (w/v) oxalic acid for 5 min.
  11. Wash for 5 min with three changes of running tap water to remove excess of 5% (w/v) oxalic acid.
  12. Place filters in the prepared protargol solution at room temperature (To prepare the protargol solution: flame 0.5 g of spiral copper wire and four copper plates of 0.1-mm thick in a Bunsen burner. Plunge in 100% methanol to expose a bright surface. Once the coil is dry place it in the bottom of the Columbia jar, add 10.0 ml of distilled water and dissolve 0.3 g of Protargol (Polysciences Inc. Cat No. 01070 or make your own- see Pan et al. 2013). Transfer filters back to back in the Columbia jar. Place one copper plate between each pair of filters.
  13. After 24 h remove the filters from the protargol solution and transfer them to hydroquinone solution (1% (w/v) hydroquinone dissolved in 5% (w/v) sodium sulfite solution and 4% (w/v) anhydrous sodium carbonate) for 5 min.
  14. Wash as in step 12 to remove excess of hydroquinone solution.
  15. Place filters in 0.5% (w/v) gold chloride for 1 min. Replace gold chloride when debris has accumulated and when the color is pale yellow.
  16. Place the filters in 2% (w/v) oxalic acid for 2 min.
  17. Wash as in step 12 to remove excess of oxalic acid.
  18. Transfer filters to 5% (w/v) sodium thiosulfate (Na2S2O3) solution for 5 min.
  19. Wash as in step 12 to remove excess of sodium thiosulfate.
  20. Dehydrate in isopropyl alcohol series (30-50-70-95-100 I-100 II-100 III). Let the filters stand at least 5 min in each alcohol. At each staining run of 8 filters maximum, discard the 100 (I) alcohol and completely dry the Columbia jar. Then, make 100 (II), the new 100 (I), and 100 (III), the new 100(II). Always fill the dry Columbia jar with fresh 100% isopropyl alcohol for a new 100 (III).
  21. Clear the filters in three changes of 100% xylene series. Change the xylenes at each run so the final change is always new as in the alcohols.
  22. Place the filters in a 20% xylene: 80% Permount ™ solution for 24 h.
  23. Mount each filter agar side up on a slide by placing a few drops of Permount below and above the filter. Cover the filter with 22 X 30 mm coverslips.

Note: Since protargol will stain both, the nucleus and the basal bodies, it allows the simultaneous assessment of both, ciliates and testate amoebae. Staining of the nucleus is important to differentiate between live and dead testate amoebae.