Soil Ecology Protocols

 

        

        E.S.S.R.E. follows a modified variation of the Smithsonian Institution's Monitoring and Assessment of Biodiversity Program protocol.  Using 20 meter by 20 meter plots divided into four quadrats, we perform a general biota survey for 22 environmental factors in the soil ecosystem.  These include:

Calcium pH Phosphorus Sulfate
Potassium Ferric Iron Magnesium Yeast
Chloride Aluminum Manganese Algae
Nitrite Nitrate Ammonia Soil Texture
Heterotrophic bacteria Mold Protozoa  
Arthropods Plant Diversity Annelids  

        We use the LaMotte STH-14 Series soil test kit to determine the various inorganic and salt levels and a standard transect analysis for plant diversity.  For the remaining items, we use the following protocols adapted from Methods for the Examination of Organismal Diversity in Soils and Sediments (CAB International, 1996) for the high school laboratory.  For inexpensive alternatives to the more costly of these protocols, see our "low-tech" serial dilutions, "low-tech" protozoa extraction, and "low-tech" arthropod procedures. Over the course of the program, protocols have been changed and improved in order to attain more accurate results. The years in which each arthropod density and diversity protocol was used is indicated below.

 

I. Estimating Soil Heterotrophic Bacteria, Yeast, & Mold Densities 

    For all three of these groups of organisms, the standard protocol is to perform a serial dilution of a soil extract in sterile water.  The step-by-step procedure for doing so is as follows:

  1. Place 1 cc of your soil sample into a culture tube containing 10 ml of sterile water; cap the tube and shake vigorously.
  2. Using a serological pipette, remove 1 ml of the soil/water mixture and place into a fresh culture tube.
  3. Add 9 ml of fresh sterile water to this second tube; cap and shake vigorously.
  4. Repeat step 2 using the second, diluted tube and then repeat step 3 with this third tube.
  5. Continue step 4 with each additional tube until you have diluted the original soil/water mixture a minimum of four times (a 10-4 dilution). You should now have a total of five culture tubes.
  6. Plate 100 l samples from the 4th and 5th tubes (dilutions 10-3 & 10-4) onto their own separate, individual petri plates filled with nutrient agar and allow to incubate at room temperature over night.
  7. Examine each of the plates for individual bacteria colonies and choose the plate with the fewest colonies to make your estimates of the number of bacteria in the original 1 cc soil sample (# colonies on plate C 102 = # of bacteria in dilution tube; # of bacteria in dilution tube C 10 |# of dilutions| = # of bacteria in original sample tube).
  8. If there are not individual colonies but still a "lawn" at the 10-4 dilution, repeat steps 1-6, adding a 5th dilution, 6th dilution, etc. as necessary until individual colonies are observed.

II. Estimating Soil Protozoa Density

  1. Place a 2 cm diameter X 15 cm core sample of soil into the bottom of a clean, empty petri dish; and allow to dry completely.
  2. Sift 9-10 g of the soil into a 2nd clean petri dish using a 1 mm2 nylon screen or mesh.
  3. Add 20 ml of distilled water to saturate the soil
  4. Cover the petri dish with its lid and allow to sit for 7 hours.
  5. Place the soil sample in a modified Uhlig extractor containing 30 ml of distilled water for 24 hours.
  6. Remove the filtrate and filter a 2nd time using 12.5 cm qualitative filter paper.
  7. Using a capillary tube, deposit 7 l of methyl-green stain on a clean microscope slide (1 l = 1 drop from the capillary tube). Then using a  disposable graduated Beral-type pipette, add 18 l (the first demarcation on the pipette) of the 2nd filtrate from step 6 to the stain on the microscope slide and cover with an 18 x 18 mm2 cover slip.
  8. Examine under a light microscope at 40X (or 100X for qualitative observations) of the various protozoa living in the soil.
  9. Use the following equation to determine the population density of protozoa in the soil sample:
  10. [(# per field of view at 40X) (total ml of water used) 747] (grams of sifted soil ) = # of protozoa per gram of soil

III. Estimating Soil Algae Density

Method used from 2001-2016

  1. Collect 36 clean glass slides, 12 white marker flags, a permanent marker, distilled water and rubber bands.
  2. On 3 of the clean slides, use a ruler and measure from the top of the slide, 1.5 cm down, and mark this point with a permanent marker.
  3. Insert longer portion of the slide into the soil until the mark is a the level of the soil, leaving 1.5 cm of the microscope slide exposed.
  4. Place a white flag next to the slide to make it easy to collect the slide at a later time.
  5. After all algae slides are in place in the soil, let slides sit in the soil for 2 days.
  6. After 48 hours, CAREFULLY remove the slide from the soil and rinse off excess particulates and debris using distilled water.
  7. Take 2 additional clean slides and place on one on either side of the algae slide. 
  8. Secure with a rubber band for purposes of carrying back to the lab.
  9. In the lab, carefully dry the outside slides and place the 3 still sandwiched together onto the stage of a compound light microscope.
  10. At 40X, count the number of algae (will have a blue-green hue) in the field of view and record.  
  11. Move the slide to a different field of view and repeat step 10.  
  12. Repeat steps 10 & 11 until at least 5 unique fields of view have been observed. 
  13. Average the 5 algae counts and divide this number by 17.3 mm2 to determine the number of algae per mm2 of soil..

Method used in 2017

  1. Collect 36 clean glass slides, 12 white marker flags, a permanent marker, distilled water and rubber bands.
  2. On 3 of the clean slides, use a ruler and measure from the top of the slide, 1.5 cm down, and mark this point with a permanent marker.
  3. Insert longer portion of the slide into the soil until the mark is at the level of the soil, leaving 1.5 cm of the microscope slide exposed.
  4. Place a white flag next to the slide to make it easy to collect the slide at a later time.
  5. After all algae slides are in place in the soil, let slides sit in the soil for 2 days.
  6. After 48 hours, CAREFULLY remove the slide from the soil and rinse off excess particulates and debris using distilled water.
  7. Take 2 additional clean slides and place on one on either side of the algae slide. 
  8. Secure with a rubber band for purposes of carrying back to the lab.
  9. In the lab, remove the sandwich slides and dip and swirl the original slide in a 500m beaker of distilled water to remove any remaining dirt.
  10. Dip one of the sandwich slides in water as well and re-sandwich the bottom of the original slide.
  11. Then, carefully dry the outside slides and place the 3 still sandwiched together onto the stage of a compound light microscope.
  12. At 40X, count the number of algae (will have a blue-green hue) in the field of view and record.  
  13. Move the slide to a different field of view and repeat step 10.  
  14. Repeat steps 10 & 11 until at least 5 unique fields of view have been observed. 
  15. 15. Average the 5 algae counts and divide this number by 17.3 mmto determine the number of algae per mm2 of soil. 

IV. Estimating Arthropod Density & Diversity

  1. Place a 1/2 meter x 1/2 meter grid on the soil you wish to survey.
  2. Carefully remove any plant material or leaf litter from within the grid and sort through it by hand, identifying and recording any arthropods found in this material (a good dichotomous key is available from the American Museum of Natural History).
  3. Next, use hand trowels to remove the soil inside the grid to a depth of 15 cm. and place the soil into a tub for sorting.
  4. Remove handfuls of the soil from the tub and sort by hand, identifying and recording any arthropods found and keeping a running tally of how many earthworms are found.
  5. Remove handfuls of the soil from the tub and sort by hand, identifying and recording any arthropods found and keeping a running tally of how many earthworms are found.
  6. To estimate the number of earthworms and each category of arthropods per square meter, multiply all final values by 4.

V. Estimating Annelid Density 

  1. Place a half meter by half meter frame onto the soil being studied and use gloves to clear all ground clutter and cover, carefully checking for any annelids.
  2. With a trowel and a ruler, excavate the 0.5 m X 0.5 m square to a depth of 15 cm, carefully placing all removed soil in a bin.
  3. Once the hole is 15 centimeters deep begin filling the hole again with the soil in the bin while checking thoroughly for any annelids and recording the number found.
  4. Once everything is sifted through, gently pat the soil down and replace the ground cover.
  5. Multiply the recorded values by 4 to determine the number of each group of annelids per square meter of soil.

VI. Soil Texture

  1. Collect a sample of soil using a trowel (the amount depends on the size of jar used; you want enough soil to fill your choice of container 3/4 full).
  2. Place the soil in a flat sided, flat bottom jar or other container where the sides and the bottom form an exact 90 degree angle (we use plastic urinalysis sample jars; they have a nice seal, are inexpensive, and reusable).
  3. Add water to the jar until about 1 cm from the top of the jar (tap or distilled water may be used).
  4. Using a pipette, add 3-5 drops of commercial dishwashing detergent (or other detergent that contains phosphates) to the soil-water mixture.
  5. Cap the jar and shake vigorously until well mixed.
  6. Let sit for several hours until completely settled.
  7. There should be 4 different layers in the jar- sand, silt, clay and water (in that order from the bottom up).
  8. The sand layer will be at the bottom of the jar, and will probably be either a lighter color than the silt layer or may be the same color, but will have a grainy texture.
  9. The silt layer will be the second to bottom and will look smooth and will have a dark brown color.
  10. The clay layer will look extremely smooth and be above the silt layer, and should be a lighter tan color.
  11. Above the clay is the remaining water from the jar.
  12. Using a ruler, measure from the bottom of the sand layer to the top of the clay layer. Record this measurement in cm.
  13. Measure the length of column of each individual layer (sand, silt, clay) and record each individual measurement in cm.
  14. Divide each individual measurement by the total measurement for the sand-silt-clay layers to find the percentage of each different type of soil in the sample.