Protocol

 

Authors

Angela Yin Yee Kong¹ and Johan Six²

Author affiliations

¹Goddard Inst. for Space Studies, 2880 Broadway, New York, NY 10025

²Department of Plant Sciences, University of California,One Shields Avenue, Davis, CA 95616

Overview

This protocol details how to collect rhizosphere soil from the field and then isolate different biophysical microenvironments, i.e., microaggregates (53-250 μm soil aggregates) versus particulate organic matter (>250 μm) and silt-and-clay particles, from both rhizosphere and non-rhizosphere soil. The rhizosphere is operationally defined as the soil fraction that adheres to root surfaces following gentle shaking of the plant root-soil system and the soil that does not adhere after shaking is the non-rhizosphere soil. From this procedure, six soil fractions are produced from one plant root-soil system: >250 μm particulate organic matter, microaggregates, and silt-and-clay fractions from rhizosphere soil and >250 μm particulate organic matter, microaggregates, and silt-and-clay fractions from non-rhizosphere soil. Additional biological and chemical analyses, such as total carbon and nitrogen content, phospholipid fatty acid (PLFA), pyrolysis, etc., can be performed on the different fractions to understand the effects of different biophysical microenvironments on soil properties and dynamics.

Background

Soil aggregate dynamics regulate carbon and nitrogen cycling within ecosystems. Aggregates not only physically protect soil organic matter (e.g., Tisdall and Oades 1982), but also influence microbial community structure (e.g., Hattori 1988), limit oxygen diffusion (e.g., Sexstone et al. 1985), regulate water flow (e.g., Prove et al. 1990), determine nutrient adsorption and desorption (e.g., Wang et al. 2001), and reduce run off and erosion (e.g., Barthes and Roose 2002). Furthermore, studies have shown the importance of microaggregates (e.g., Jastrow, 1996; Six et al., 1998; Gale et al., 2000) and especially microaggregates-within-macroaggregates (Six et al., 2000; Denef et al., 2004; Kong et al. 2005) in the protection and stabilization of carbon and nitrogen.

The dynamic nature of the rhizosphere-soil structure complex createsa mosaic of physicochemically varying microenvironmentsin the soil. Microorganisms preferentially colonize the rhizosphere because root exudates (i.e., sloughed cells, secretions and exudates) are a major source of nutrients in soils, making the rhizosphere an area of intense activity with specific biological, chemical, and physical characteristics(Lynch and Whipps, 1990; Kennedy, 1998). Microaggregates formed in the rhizosphere are stabilized by intimate contact with clay, organic materials, and amorphous inorganic components (Turchenek and Oades 1976). Outside the rhizosphere, microaggregate formation is influenced by i) microbially-produced polysaccharides which bind minerals and ii) the direct interaction of organic materials with inorganic colloids.

Materials/Equipment

Collecting Rhizosphere Soil

• Equipment to extract plant root-soil system from the field (e.g., soil cores or shovels)
• Separate containers to store rhizosphere and non-rhizosphere soil samples
• Large bin

Isolating Microaggregates from Rhizosphere Soil

• Microaggregate isolator (see figures below)
• Reciprocal shaker to mount the microaggregate isolator
• 4 mm diameter glass or stainless steel beads
• Pre-weighed and labeled specimen cups
• 53 μm sieve
• 250 μm sieve
• small rubber stopper that fits into the funnel of the microaggregate isolator
• Two or more 2 L beakers
• Flocculant (e.g., 0.25M CaCl₂, 0.25M MgCl₂, or 12M HCl)
• 5000 L pipetman
• Squirt water bottles
• 250 or 760 mL centrifuge bottles

Units, terms, definitions

• DI water = deionized water
• m = micrometer

Procedure

Collecting Rhizosphere Soil

In the field

1. Identify desired plant root-soil system in the field and extract using method that is appropriate for your study and the environmental conditions (henceforth, this sample of the plant root-soil system will be referred to as the -soil core’).

• For example, in a maize-tomato cropping system, an herbaceous winter cover crop soil core was extracted by inserting a 30 cm diameter polyvinylchloride (PVC) core to a depth of 30 cm below the cover crop shoot-soil-surface, and then the entire core containing the root-soil system was extracted.

2. Ideally, store soil cores at 4^∘C until they are separated into rhizosphere and non-rhizosphere soil.

• The purpose of storing at 4^∘C is to maintain field moisture in the soil core, while limiting microbial activity and, therefore, decomposition and changes to aggregate structures in the soil core see Notes below; however, lengthy storage periods at 4^∘C should be avoided.

In the laboratory

3. Remove soil core from 4^∘C storage.

4. Divide the soil core into smaller subsections that run from the soil surface to the bottom of the core.

• While a few subsections can be left at room temperature before proceeding to Step #3, subsections that will not be processed within 20 minutes should be returned to 4^∘C (i.e., three or four subsections can be processed at room temperature within 20 minutes).

5. Working one subsection at a time, grip the exposed part of the plant shoots and then shake the mass of roots and soil over a large bin.

• The soil still adhering to the roots after no more soil can be separated from the root due to the shaking is the rhizosphere soil and this can be placed in a container.
• Soil that did not adhere to roots is the non-rhizosphere soil and can be removed from the bin and placed in a container separate from the rhizosphere soil sample.
• Most likely, there will be substantially more non-rhizosphere soil than rhizosphere soil.
• Although the amount of time and force necessary to shake the non-rhizosphere soil from the root mass-soil sample will depend on several factors (e.g., root architecture, soil type, soil moisture content, etc.), the duration of the shaking should be on the order of a few seconds and not likely more than 30 seconds.

6. Repeat Steps #3-5 for remaining subsections of a soil core.

7. If a quantification of soil moisture is desired, then take a subsample from each rhizosphere and non-rhizosphere soil sample and proceed with chosen soil moisture measurement procedure.

8. Once collected, store rhizosphere and non-rhizosphere samples at -20^∘C until the physical fractionation process.

• Storage at -20^∘C will greatly reduce degradation of the root, but if the operator is prepared to proceed to the next steps within 24 hours of collection, then storage at 4^∘C is likely adequate.

Isolating Microaggregates from Rhizosphere Soil

Microaggregate isolator assembly; Please refer to Figure 1, -Microaggregate Isolator Components Diagram’ and images (Figs. 2-6) below for a visual guide to the assembly of parts and connections

9. Assemble microaggregate isolator:

• Assemble the Column Assembly (lid (Figure 2; Part #1 in Figure 1) + column (Figure 2; Part #2 in Figure 1) + 250 μm screen (Figure 3; Part #3 in Figure 1))
• Assemble the Funnel Assembly (funnel (Figure 4; Part #4 in Figure 1) + stainless steel backing (Part #5 in Figure 1))
• Using six screws (Figure 4), join the Column Assembly to the Funnel Assembly (Figure 5).

10. Affix stainless steel backing of microaggregate isolator assembly onto reciprocal shaker with eyelet screws.

11. Connect the 5/8 inch diameter piece of Tygon tubing to the funnel of the isolator, clamp the tubing to a stand and make sure that the height of the arc of the tubing is level with a height 2 cm above the 250 μm sieve; Rest the end of the 5/8 inch Tygon tubing on the 53 mm sieve + receiver pan (53 m sieve assembly; Figure 6).

• The size of the receiver pan will depend on the diameter of the 53 μm sieve used and amount of water generated during the average microaggregate isolation procedure.
• It is suggested that a splash guard (e.g., 10cm wide strip of aluminum foil) be placed around the rim of the 53 μm sieve, at the end of the 5/8 inch tubing, so that the material coming from the tubing will not splash out of the 53 μm sieve.

12. Plumbing:

• Connect one end of ¼ inch tubing to DI water source and other end to one opening of the Y-connector (Part #7).
• Onto another opening of the Y-connector, connect another length of ¼ inch tubing; at the end of this tubing, connect the smaller regulator valve (Part # 8) that has another length of ¼ inch tubing at the opposite end of the valve.
• Connect a piece of ¼ inch tubing to the last opening of the Y-connector; connect the end of this tubing, to the end of the larger valve (Part #6) with an open-close knob; then, connect the other end of the valve to the column lid with a piece of ¼ inch tubing.

13. Turn on DI water and start flow of water into the isolator unit by turning the open-close knob on the valve (Part #6). If necessary, increase or decrease the water flow in order to have a slow but stable water flow through the isolator, by adjusting the knob on Part #6 or Part #8 (it is not advised to change the flow of water at the DI water faucet).

14. Fill isolator with DI water to 2 cm above the 250 μm sieve screen in the column assembly. The water level should remain at this level and the excess water should be flowing from the 5/8 inch Tygon tubing into the 53 μm sieve assembly. Check to see if the water level remains 2 cm above the 250 μm sieve screen by closing the valve (Part #6) to the DI water source.

15. With the valve closed (Part #6), add 50 glass or stainless steel beads (4 mm diameter) into the column/atop the 250 μm screen, resume the flow of water, turn on the reciprocal shaker to a low speed for several seconds, then shut off the reciprocal shaker. Check to see that the 2 cm head is stable/unaffected.

16. Once the 2 cm head and stable water flow has been achieved, cut off the water supply to the isolator by closing the valve (Part #6). Pour out the water collected in the receiver pan and immediately replace the 53 μm sieve assembly beneath the 5/8 inch Tygon tubing.

Microaggregate isolation from rhizosphere soil

17. Remove and thaw frozen rhizosphere and non-rhizosphere samples from -20^∘C storage (~10 minutes).

18. Number (sequentially) and record weights of at least 7 specimen cups per sample (i.e., label with # and record weight on the cup).

19. Add ~20g non-rhizosphere or ~25-30g rhizosphere soil subsamples into the microaggregate isolator column and place the lid on the column assembly.

20. Start water flow through the isolator by opening valve on Part #6 and turn on the shaker to a low speed (~150 rpm). Shake the device until water flowing out of the device onto the 53 μm sieve is clear and all aggregates on top of the 250 μm screen are broken up (check by removing the lid and looking into the column).

• The length of shaking time necessary can vary from 3 to 20 minutes (depending on various factors mentioned in the Notes below); a large quantity of water plus silt-and-clay may collect in the receiver pan underneath the 53 μm sieve after 5 minutes of shaking, so be careful for spills.
• You may want to stop water flow to the isolator (close valve in Part #6) in order to see if all the aggregates are broken up, but DO NOT turn off the faucet or else air will be caught in the tubes.

21. Stop the water flow to the isolator (closing Part #6) and turn off the DI water faucet.

22. Rinse off the sides of the column with DI water in a squirt bottle.

23. Drain the 5/8″ tubing completely onto the 53 μm sieve by unscrewing the eyelet screws and removing isolator from shaker and raising the isolator above the 53 μm sieve. This should eject most of the <250 μm material quickly onto the 53 μm sieve.

24. Re-attach the isolator onto the shaker, remove the 5/8″ tubing from the end of the funnel, plug end of the funnel with rubber stopper and then rinse 5/8″ tubing with DI water onto the 53 μm sieve and make sure the tubing is completely clean.

25. Manually sieve the material left on the 53 μm sieve for two minutes by moving the sieve 50 times, in an up-and-down motion, with a slight angle to ensure that water and small particles go through the mesh (Elliott, 1986; see link to video protocol in -Other resources’ below).

26. Transfer water and particles that went through the 53 μm sieve into a 2 L beaker (if not processing biophysical microenvironments for biological variables, then see Notes below).

27. Calculate amount of flocculant to add to <53 μm material to achieve a theoretical concentration of 0.005M HCl (i.e., treat water+silt+clay solution as pure water)

• For example, for a 2 L silt-and-clay solution, add 833 L HCl.
• 0.25M MgCl₂ + 0.25M CaCl₂ can also be used as flocculants.

28. Add appropriate amount of flocculant to the sample with a pipetman and let solution stand/flocculate for 15 minutes.

29. Meanwhile, unscrew eyelet screws and remove isolator from shaker.

30. Remove rubber stopper and using a squeeze bottle with DI water or similar set-up, run water through device to rinse material and beads onto a 250 μm sieve and transfer this material to a pre-weighed specimen cup for the coarse particulate organic matter (>250 μm material).

31. Again, using a squeeze bottle with DI water, transfer the microaggregates on the 53 μm sieve into a pre-weighed specimen cup.

32. After 15 min of flocculation, transfer the silt-and-clay solution into centrifuge bottles (using as little DI water as possible).

33. Balance the centrifuge bottles and centrifuge the samples at 5,000 RPM (or equivalent RCF) at 4^∘C for 15 minutes.

34. Decant the supernatant and consolidate the pellets into fewer bottles.

35. Centrifuge these samples once more at 5,000 RPM at 4^∘C for 15 minutes.

36. Collect silt-and-clay pellets into pre-weighed specimen cups.

37. Store all samples in specimen cups in -20^∘C freezer until further analysis.

Figures

Figure 1. Microaggregate isolator components.

Figure 2. Column Assembly – lid and column (Parts #1 and #2, respectively,in Figure 1).

Figure 3. Column Assembly – 250 μm screen (Part #3 in Figure 1). Placement of 250 μm screen is between the Column Assembly and the Funnel Assembly.

Figure 4. Funnel Assembly – funnel (Part #4 in Figure 1) and location of screws necessary to attach Column Assembly to Funnel Assembly.

Figure 5. Column Assembly joined to the Funnel Assembly.

Figure 6. Column Assembly plus Funnel Assembly connected via 5/8 inch Tygon tubing to the 53 m sieve assembly (53 μm sieve + receiver pan). Note that lid of the Column Assembly shown here is a variation of the lid shown in Figure 2.

Other resources

– Instructional video of sieving procedure used to separate microaggregates from the silt-and-clay particles after microaggregate isolation:

Soil fractionation (produced by J. Denbow and J. Six).

– A video protocol for the complete microaggregate isolation procedure can be found here:

Microaggregate isolation (produced by J. Denbow, J. Sheehy, and J. Six)

Notes and troubleshooting tips

Collecting Rhizosphere Soil

• After the extraction of the plant root-soil system, it is best to process the core as soon as possible for two reasons: 1) to maintain field moisture in the core, as collecting rhizosphere soil from the core is easier with field moist soil than from a core that has dried (the rate of drying after extraction from the field will vary by soil type and ambient temperature) and 2) limit biological activity in the core (storing at 4^∘C will help with this to some degree).
• It is impossible to remove roots from the non-rhizosphere soil completely. It is important to attain consistency in the visual criteria for rhizosphere soil to minimize core-to-core variation. The time necessary to collect rhizosphere soil will depend on the size of the core, the root architecture of the plant, the age of the plant, and the soil type; for a 30 cm diameter x 30 cm deep core taken from a mature herbaceous plant root-loamy soil system, the procedure took ~45 minutes per core.
• Both the procedures for the collection of the rhizosphere soil and the isolation of the microaggregates are operator-sensitive, but need to be standardized across samples that are being compared.

Isolating Microaggregates from Rhizosphere Soil

• During the microaggregate isolation (Step #20), watch the water level within the column during shaking as the 250 μm screen can become clogged or air pockets can form in the 5/8 inch tubing and can lead to accumulation of the soil solution greater than a 2 cm height in the column.
• The length of shaking time necessary to completely isolate microaggregates from the soil sample varies depending on the stability of the aggregates, the size of the soil sample, the flow rate of the DI water, and the type of 4 mm diameter beads used (glass or stainless steel). For example, isolation of microaggregates from 10 g of a grassland soil, using stainless steel beads, can take nearly 20 minutes.
• If the biophysical microenvironments will not be analyzed for biological variables after Step #25, then transfer the material on the 250 mm screen and the 53 mm sieve, and the silt-and-clay solution into tared aluminum pans via backwashing (see Fig. 7 below). The microenvironments can then be dried at 60^∘C and analyzed (e.g., for total carbon or nitrogen).

Figure 7. Backflushing material from sieves with DI water in squirt bottles into tared aluminum pans.

• This protocol has been successfully coupled with stable isotope probing and compound specific stable carbon isotope analysis of microbial phospholipids fatty acids (PLFA) to directly link microbial community structure and activity within different cover crop-enhanced soil microenvironments (Kong et al. 2010, 2011; Kong and Six, in prep).

Other:

• Species on which this protocol has been used: Vicia dasycarpa, hairy vetch (an herbaceous plant)
• Setting in which protocol has been tested: Field (temperate cropping system) and laboratory

Suppliers

• Centrifuge: Sorvall RC-5C Plus Superspeed centrifuge, Thermo Scientific
• Reciprocal shaker: Eberbach shaker, model 6000

Literature references

Barthes B, Roose E (2002) Aggregate stability as an indicator of soil susceptibility to runoff and erosion: validation at several levels. Catena 47, 133-149.

Denef K, Six J, Bossuyt H, Frey SD, Elliott ET, Merckx R, Paustian K. (2001a) Influence of wet-dry cycles on the interrelationship between aggregate, particulate organic matter, and microbial community dynamics. Soil Biol. Biochem. 33, 1599-1611.

Elliott ET (1986) Aggregate structure and carbon, nitrogen, and phosphorus in native and cultivated soils. Soil Sci. Soc. Am. J. 50, 627-633.

Gale WJ, Cambardella CA, Bailey TB (2000) Root-derived carbon and the formation and stabilization of aggregates. Soil Sci. Soc. Am. J. 64, 201-207.

Hattori T (1988) Soil aggregates in microhabitats of microorganisms. Rep. Inst. Agr. Res. Tohoku Univ 37, 23-36.

Jastrow JD (1996) Soil aggregate formation and the accrual of particulate and mineral-associated organic matter. Soil Biol. Biochem. 28, 665-676.

Kennedy AC (1998) The rhizosphere and spermosphere. In: Sylvia DM, Fuhrmann JJ, Hartel PG, Zuberer DA (eds.). Principles and applications of soil microbiology. Upper Saddle River, New Jersey, Prentice Hall. pp. 389-407.

Kong AYY, Six J. Cover crop root-C assimilation into soil microbial communities within soil microenvironments of alternative and conventional cropping systems. In prep.

Kong AYY,Hristova K, Scow KM, Six J (2010) Impacts of different N management regimes on nitrifier and denitrifier communities and N cycling in soil microenvironments. Soil Biol. Biochem.42, 1523-1533.

Kong AYY, Scow KM, Córdova-Kreylos AL, Holmes WE, Six J (2011) Microbial community composition and carbon cycling within soil microenvironments of conventional, low-input, and organic cropping systems. Soil Biol. Biochem. 43, 20-30.

Lynch JM, Whipps JM (1990) Substrate flow in therhizosphere. Plant Soil 129, 1-10.

Prove BG, Loch RJ, Foley JL, Anderson VJ, Younger DR (1990) Improvements in aggregation and infiltration characteristics of a krasnozem under maize with direct drill and stubble retention. Aust. J. Soil Res. 28, 577-90.

Ranjard L, Richaume AS (2001) Quantitative and qualitative microscale distribution of bacteria in soil. Res. Microbiol. 152, 707-716.

Tisdall JM, Oades JM (1982) Organic matter and water-stable aggregates in soils. J. Soil Sci. 62, 141-163.

Turchenek LW, Oades JM (1978) Organo-mineral particles in soils. p.137-144. In Emerson WW et al. (eds.). Modification of Soil Structure. Wiley, Chichester.

Sexstone AJ, Revsbech NP, Tiedje JM (1985) Direct measurement of oxygen profiles and denitrification rates in soil aggregates. Soil Sci. Soc. Am. J. 49, 645-651.

Six J, Elliott ET, Paustian K (2000) Soil macroaggregate turnover and microaggregate formation: A mechanism for C sequestration under no-tillage agriculture. Soil Biol. Biochem. 32, 2099-2103.

Wang X, Yost RS, Linquist BA (2001) Soil aggregate size affects phosphorus desorption from highly weathered soils and plant growth. Soil Sci. Soc. Am. J. 65, 139-146.

Health, safety & hazardous waste disposal considerations

• 12M HCl is a very strong acid; take proper precautions outlined by your institute’s health and safety department when using this chemical