Soil Sampling

From livingsoilsp2011.wordpress.com/student-support/sampling:

• Microscope: Binocular 10 x W.F. eyepeice, 4x lens, 10x lens, 40 x lens, ABBE condenser, N.A. 1.25, iris diaphragm
• Microscope cover
• Microscope case if moving frequently
• Computer for spread sheet
• 1/4 tsp
• Glass vile with lid (3) or medicine spoons (3) with mL increments
• Glass slides
• Glass cover slips – 22 X 22 mm(preferred) or 22 X 18 mm or 18 X 18 mm
• Lens cleaning paper
• Eye dropper and holder or pipet
• Towel or paper towels
• Spring water: 1 gal
• Squirter bottle: for spring water to make dilutions
• 2 Empty vessels: to squirt out excess liquid from rinsing eye dropper
• Access to hand wash station with soap: sometimes the material you test can be pathogenic

• Solute: Material to be diluted
• Solvent: Material that is diluting
• Dilution Factor: Ratio of final volume
• 1:5 Dilution = 1 unit volume solute + 4 unit volumes solvent = dilution factor of 5

A series of simple dilutions which amplifies the dilution factor

• The source of the dilution material of the previous step
• Total dilution factor: The product of the individual dilution factors in each step up to it.

You may find that a 1:5 dilution is appropriate for compost but not soil; soil likely requires making serial dilutions (see Making a Serial Dilution below)

• Make a simple dilution (1:5 dilution)
• Measure 1/4 tsp solute (material to be diluted – soil or compost) put this into the glass vile.
• Measure 4 mL of solvent (material that is diluting – distilled water) in the medicine spoon and then put it into the glass vile with the solute and put the lid on the vile.
• Break up the aggregates in the 1:5 dilution by shaking the vile for 30 seconds (count one one thousand, two one thousand...) Have a cadence and keep your elbow raised up 90º to your torso, with your hand going to your shoulder and then down so that your arm is horizontal to the ground. Vigorous shaking can injure the critters so don’t be too rough

Material does not need to be diluted. If you are doing a qualitative analysis then you may need to dilute this material to do bacterial counts. This is the same procedure as above, start with a simple dilution and proceed with serial dilutions as needed (see Making a Serial Dilution below).

• Clean slide and cover slip with lens cleaning paper so that both are perfectly clean
• Use eye dropper and squeeze three times into the vile with dilution to mix it up
• Place appropriate amount of drops on your slide. In general: 1 drop for 20 X 18 and 18 X 18, 2 drops for 22 X 22
• Squirt Excess water back into the glass vile with dilution in it
• Before placing the dropper back into the holder, submerge into the vessel with the spring water and suck up water, squirt the water out into the empty vessel. Do this two more times
• Place cleaned eye dropper into the holder and suck up water into the eye dropper. Do this every time to keep the holder clean, they are hard to clean AND you don’t want it contaminated because it will alter the results of your assay
• Hold cover slip by the edges to keep it clean, place the edge of the coverslip on the slide and drag it gingerly across the drop to spread it out
• Place cover slip on slide: there should be zero air bubbles and zero liquid oozing out on the slide
• If you cannot see through the material on the slide, it’s too dense, now it’s time to make a serial dilution

• Starting from a 1:5 dilution, add 5 mL distilled water to the vile. Mix for 30 seconds as you did with the 1:5. You now have a 1:10 dilution
• Fill the medicine spoon with the 1:10 dilution (use the dropper) up to 1 mL
• If you add 1 ml you will have a 1:20, adding 2 mL = 1:30, 3mL = 1:40, 4 mL = 1:50, if you add 9 mL it will be a 1:100 dilution
• Take 1 mL from the 1:100 dilution and place it into a new medicine spoon. Adding 4 mL will give you a 1:500 dilution, adding 9 mL will = 1:1000 dilution

Note: Everything increases by a factor of 10

Pick a corner on the slide and scan the entire slide at 40X TM (total magnification) this means looking through the 10x eye piece while using the 4X lens. If you find that you cannot identify anything because there is too much material in the way, add 5 mL of water to your vile (you now have a 1:10 dilution). If you still can’t see, continue making serial dilutions until you can. Keep track of the dilutions though.

Scan for nematodes, to identify type zoom in up to 400x TM (using the 40x lens) record findings appropriately.

Make note of:

• OM present
• Acids (fulvic/humic) present or not
• Microarthropods
• Larvae
• Aggregation

Use to hone in on objects and bring them into focus before going up to the 40X lens

Done at the same dilution as when scanning for nematodes or if needed go up

Look for:

• Protozoa
• Fungi
• Actinobacteria
• Make note of diversity in bacteria

Other indicators of + or – such as anaerobic indicators, bad fungi, bacterial clumping...

• Use medicine spoon to determine how many drops it takes for you individual dropper to fill 1 mL of water
• In Columns Y13, Y18, Y21, Y28, Y35, Y36, Y37, Y41 you will have to change the value for the number of fields of view on your slide as well as the number of drops it takes to fill 1 mL with your dropper AND that number will depend on your placing 1 drop or 2 on the slide and the size of cover slip you are using
• If it takes 20 drops to fill 1 mL and I place one drop on my slide, the value entered will be 20 drops. However, if I use 2 drops on the slide the value will be 10 drops
• Change the dilution factor as appropriate
• Is the standard deviation at or below < 30% of the mean?
• (mean)(0.3) = 30%. Is this at or below the standard deviation? Stop at 5 FOV. If not, keep going to 10 FOV or recount the outlying count

• Testing for your dropper
• Changing the factors in column Y
• Dilution rates
• Standard deviation is < 30% of the mean in the bacterial category
• Bacterial diversity notes
• Observations: humic acid presence, fulvic acid presence, aggregation developement, organic matter presence
• Summarise: F:B ratio and where it fits into succession. What needs to be done to create a more productive environment. Are we meeting minimum requirements for soil food web presence?