|CSS 360-Soil Biology
9 November 2011; Lab
Microfauna: Protozoa, Nematoda, Tardigrada and Rotifera
We will learn methods for extracting protozoans, nematodes, tardigrades and rotifers from soil.
Teaching lab PSSB
Soil microfauna require water films/water filled pore spaces for feeding, reproduction and dispersal. Accordingly, techniques designed to assess the abundance and diversity of microfauna take advantage of the dependence of these organisms on water. We will use a plate culturing method for the assessment of protozoans and Baermann funnels to extract nematodes, rotifers and tardigrades from field-collected soil from 2 ecosystems – a fallow field and a deciduous forest.
Materials – Each group will need:
one 24well plate (all wells are lined with soil-amended agar)
125ml specimen cups (6)
1000 uL pipet tips
1000 uL pipettor
Oak Ridge centrifuge tubes (6)
Plastic weight boats (6)
1. Weigh 10g of each of your two soils three times and place into a 125ml specimen cup and record the soil mass. You should have 6 specimen cups, each with 10g soil, 3 from forest and 3 from field.
2. Add 40ml of DI water (1:5 dilution) to each.
3. All samples should then be placed on benchtop shaker for 15 minutes.
4. Using a pipettor and graduated cylinder measure a 5ml subsample from each 125ml bottle and transfer into a pre-labeled 15ml centrifuge tube along with 5ml of DI water (1:10 dilution). You should have 6-15 mL centrifuge tubes.
5. Using a pipettor, pipet 500 uL (0.5 mL) of the 1:10 dilution into each well of a full column on a microplate (4 wells). The 4 wells in a column represent 4 lab replicates. Each plate will hold 6 samples and each group will be using one full plate (3 replicates of forest soil, and 3 replicates of field soil).
6. Each group should take notes and keep track of sample layout within plates. Sample information should include your group # and date (ex. Grp2-11/09/11). Once all samples are pipetted, plates will be incubated at 27°C for a week. The day before our next lab I will be adding a bacterial broth to each well in order to stimulate protozoan feeding.
-Nematoda, Tardigrada, Rotifera-
Materials - Each group will need:
15ml centrifuge tubes (6)
Baermann funnels (6)
Mesh disks (6)
squirt bottle with tap water
1. Extractor preparation – Each group will have 6 Baermann funnels to set up. Funnels will be placed into rack. Binder clips will be attached to rubber hoses on stem of funnel. Each group will fill 6 funnels half way with water.
2. Place a mesh disks into each funnel. Place a drain pan under your funnel stems and lightly squeeze binder clips to get rid of any O2 bubbles trapped in the bottom of the funnel stem.
3. Sample preparation – Wearing gloves weigh 10 g of soil onto the center of a kimwipe. The mass will be recorded and the kimwipe will be wrapped into a pouch around soil. Each pouch will have a label lightly attached so that it can be transferred to a vial later. Labels should include your group #, date and habitat from which the sample was taken (ex. Grp 1-field3-11/09/11). Labels will be transferred from kimwipes to funnel stem. Kimwipe pouches are placed onto the screen mat in each funnel. Once all samples are placed into funnels, water is added to cover pouch. Saran wrap is used to cover funnels to prevent desiccation. Samples will be allowed to extract until Friday afternoon.
4. Wet:Dry Correction - Five grams of soil will be weighed and placed into aluminum tins. Exclude root material and stones from soil samples as these can strongly influence wet:dry estimates. Weight will be recorded and sample will be placed into a drying oven at 60°C for wet:dry correction.
On Friday, I will drain 7ml of water into a 15ml centrifuge tube. The tubes will be topped off with 7ml of 4% formalin to preserve the fauna.