Materials Included In Kit

Additional Materials Required

Prelab Preparation

1. Cut the white veil material into 30, 15 x 15 cm squares before student use.
2. Cut the 9" x 9" black nylon mesh squares into 15, 12 x 12 cm squares before student use.

Safety Precautions

When working with protozoa, it is important to minimize contamination of materials. Therefore, always be sure you are using carefully cleaned or sterilized Petri dishes, nylon meshes, microscope slides, cover slips, Uhlig extractors and pipets. Also, never reuse a pipet used to make a microscope slide. The protozoa themselves are harmless; so there are no real dangers with working with them. Be sure to wash hands after working with the soil samples. Wear chemical splash goggles.

Disposal

Please consult your current Flinn Scientific Catalog/Reference Manual for general guidelines and specific procedures, and review all federal, state and local regulations that may apply, before proceeding. Soil samples should be disposed of according to Flinn Suggested Disposal Method #26a. All the nylon mesh, Petri dishes, funnels, and microscope slides can be reused without any autoclaving as long as students have cleaned them thoroughly. Distilled water used in this activity does not need to be sterilized.

Lab Hints

• Collect soil samples from the top 15 cm of the location being investigated. Place and store each sample in its own separate clean plastic sandwich or freezer bag (do not reuse any bag to avoid contamination). A 1" Soil Sampling Tube is available from Flinn Scientific and includes 20 soil sampling bags (Catalog No. AB1148).
• It is very important that students collect all the samples they want to study on the same day at the same time because the soil is still going to be “alive” in the plastic bag. Only by collecting samples concurrently will you control for changes in the environment (such as a heavy downpour between one class session and the next).
• Another important point is understanding that this protocol is a procedure for estimating population levels. There is no possible way to know the exact number of protozoa in a gram of soil. Students need to keep this fact in mind when they are analyzing their data.
• The methyl green solution should be place in the pipet tips and dispensed dropwise by the release of pressure using a finger over the top of the pipet tip.
• The number 747 given in the equation in step 19 of the Procedure represents the factor needed to convert the portion of the 25 µL (stain and filtrate) of liquid being observed in a single field of view at 40X on a slide with a 18 x 18 mm cover slip to 1 mL of sample.

Teacher Tips

• This kit contains enough material for 15 groups of students working in pairs. Several days or class periods are required to complete this activity.
• There currently are no standardized keys for soil microbe identification. Consider having students develop their own system using objective qualities like colony color for identifying different bacterial or mold strains.
• Because microbe population levels are estimates and because the quantity of microbes in the soil is so enormous, contamination is never really a true problem with these protocols. Strict sterile procedure in this experiment is not necessary, and the only materials that students need to dispose of after using them are the transfer pipets.
• The modified Foissner/Uhlig protocol the students use has them counting the protozoa at the 40X power (and all the mathematical formula constants assume this as well as the size of the cover slips, amount of stain, and sample size–modification of any of these variables will change the formula). At 40X, the protozoa are still quite small and will basically appear as the large, light-blue “dots” in the field of view. You will need to help students understand that they are only counting the blue objects. One of the reasons for using the methyl-green stain is that only the protozoa turn blue; all other microbes are a different color.
• To increase accuracy, we recommend that students count more than one field of view (three minimum; five maximum) and use the average count to compute the population estimate. Students may take a more qualitative approach to studying protozoa (e.g., a diversity study), by examining their samples at the 100X or 400X power where only the blue-stained cells are the protozoa and their distinguishing features are readily seen.

Sample Data

Protozoa Data Table

References

Special thanks to David Brock, Kate Brockmeyer, Kalyani Ravi and Leah Miller for sharing this activity with Flinn Scientific.

This lesson has its origins in a curriculum developed and supported by monies from the following institutions and programs:

• Institute for Ecosystem Studies
• Paul F, Brandwein Institute
• ReliaStar/Northern Life “Unsung Heroes” Program
• Toshiba America Foundation
• Captain Planet Foundation, Inc.
• Gustav Ohaus Awards
• Waksman Foundation for Microbiology

Anderson, O. Roger & Druger. Marvin. (1997) Explore the World Using Protozoa. Washington, D.C.: NSTA Press.

Bramble, Judith E. (1995) Field Methods in Ecological Investigation for Secondary Science Teachers. St. Louis: Missouri Botanical Garden.

Cothron, Julia H.; Giese, Ronald N. & Rezba, Richard J. (2000) Students and Research: Practical Strategies for Science Classrooms and Competitions, 3rd ed. Dubuque: Kendall/Hunt Publishing Company.

Environmental Science Summer Research Experience for Young Women (2003). Available online http://faculty.rpcs.org/brockda/essre.htm

Hall, Geoffrey S., ed. (1996) Methods for the Examination of Organismal Diversity in Soils and Sediments. Paris: CAB INTERNATIONAL.

Nardi, James B. (2003) World Beneath Our Feet: A Guide to Life in the Soil. New York: Oxford University Press.

Samuels, Myra L. (1989) Statistics for the Life Science. Englewood Cliffs, NJ: Prentice Hall.

Introduction

Extract microorganisms from the soil and explore the exciting world of soil protozoa in this hands-on activity.

Concepts

• Protozoa
• Population
• Predator–prey cycles
• Habitat

Background

One of the last frontiers in science today is, ironically, the very dirt beneath our feet. Soil is literally the foundation of every terrestrial ecosystem and biome on Earth, and yet we know more about the moon than we do this realm of the common earthworm. The organisms that inhabit the soil are responsible for all of the ecological cycles that make life possible for plants and animals, but most of these microbes are poorly understood and the majority remain unidentified and unknown.

An example of an entire category of such organisms waiting to be explored are the protozoans. Because soil is seldom 100% dry, water coats the microscopic particles and pores that make up the texture of dirt, and in this very thin sheet of moisture lives a veritable zoo of these single-celled eukaryotes. The lions and tigers of their world, protozoa prey on the bacteria, decomposers and fungi that abound in soil. Just like their much larger animal “cousins,” they control population numbers and, hence, regulate the cycling of organic nutrients in their ecosystem. A few types are actively involved with nitrogen fixation, and some help protect plants, nematodes and earthworms from various soil pathogens. A few are even photosynthetic and contribute to the primary productivity in the soil. However, the majority just roam the soil’s “rivers,” “lakes” and “streams,” hunting their unwary prey.

There are more than 300 species of soil protozoa, ranging in size from 5 to 160 μm, and like other soil microbes, their quantity in any given patch of dirt is enormous—often 90,000+ in a single gram. In this activity, a modified Foissner/Uhlig procedure will be used to extract protozoa from soil and dilute the protozoa to a point where they can be counted and seen individually.

Materials

Safety Precautions

When working with protozoa, it is important to minimize contamination of materials. Therefore, always be sure you are using carefully cleaned or sterilized Petri dishes, nylon meshes, microscope slides, cover slips, Uhlig extractors and pipets. Also, never reuse a pipet used to make a microscope slide. The protozoa themselves are harmless; so there are no real dangers with working with them. Be sure to wash hands after working with the soil samples. Wear chemical splash goggles.

Procedure

Prepration—Making a Uhlig Extractor

1. Obtain one of the two large cups (cup A) and cut the bottom off the cup, up to and including the first indentation on the side of the cup, as shown in Figure 1.

2. Take the other large cup (cup B) and cut out the circle at the bottom of the cup. Then cut and make “legs” for the cup as instructed in Figure 1.
3. Turn cup A upside down and layer both pieces of the white nylon squares together over the cup opening, making sure you align the nylon mesh so that the lines of it do not cross each other.
4. Holding the white nylon square(s) in place, insert cup A into cup B as shown in Figure 2.

Experiment

1. Place a 1" diameter x 6"-long cylinder of soil into the bottom of a clean, empty Petri dish.
2. Allow to dry completely (usually 24 hours).
3. Transfer the soil to a small cup.
4. Cover the top of the cup with a square piece of black nylon mesh.
5. Use a rubber band to secure the mesh in place (see Figure 3).

6. Sift 9–10 g of the soil through the mesh into a second clean Petri dish (see Figure 4). Use a balance to measure the mass amount of the sifted soil sample in grams. Record the mass in the data table.

7. Add 20 mL of distilled water to the sifted soil.
8. Cover the Petri dish with its lid and allow to sit until the next day.
9. Place 30 mL of distilled water into a clean, empty Petri dish (the washed and dried Petri dish from step 1 may be used).
10. Set the Uhlig extractor upright in the Petri dish filled water (see Figure 5).

11. Scoop the rehydrated soil from step 7 into the bottom of the Uhlig extractor and allow it to sit for 24 hours.
12. Remove the Uhlig extractor from the Petri dish and set aside to clean according to the instructor.
13. Set up a ring stand with a ring clamp and a funnel (see Figure 6).

14. Fold a piece of filter paper and place in the funnel.
15. Pour the water from the Petri dish that contained the Uhlig extractor into the funnel and collect the filtrate in a clean, dry beaker. This filtrate contains the protozoa that will be examined under the microscope.
16. Using a yellow pipet tip, place 7 µL of methyl green stain on a clean microscope slide (7 µL = 2 drops from the pipet tip).
17. Then using the graduated pipet, add 18 µL (the first line on the graduated pipet) of the protozoa filtrate from the beaker to the stain and cover with a cover slip.
18. Examine the slide under the microscope on the 40X power and count the number of protozoa observed (they will be the blue objects). Record this number in the Protozoa Data Table. Search multiple areas of the slide and obtain four more protozoa counts. Record the counts and the average number of protozoa in the field of view in the Protozoa Data Table.
19. Use the following equation to determine the population density of protozoa in the soil sample:

Student Worksheet PDF

10747_Student1.pdf