Soil-Test Biological Activity Standard Operating Protocol 2018-02

Alan Franzluebbers USDA-Agricultural Research Service 3218 Williams Hall NCSU Campus Box 7620 Raleigh NC 27695 Tel: 919-515-1973 Email: [email protected]

Purpose The flush of CO2 following rewetting of dried soil during 3 days of incubation is a general indicator of soil heterotrophic activity [3, 10, 11]. It also relates closely to soil microbial biomass carbon and potentially mineralizable nitrogen.

Materials a. b. c. d. e. f. g. h. i. j.

60-mL graduated wide-mouth (3-cm diam) glass containers with etched lab ID number – to contain soil 1 M sodium hydroxide (NaOH) – dissolve 40 g NaOH in d.i. H2O, dilute to 1 L – to trap CO2 30-mL Nalgene screw-top HDPE vials plus screw-cap – to contain alkali Auto-dispenser – to accurately dispense 10 mL alkali 7-dram plastic vial – to contain water for humidity 1-L mason jars with rubber-seal lids – as incubation vessel Wooden dowel – to press soil to desired density 10-mL adjustable hand pipette – to dispense water to soil Incubation cabinet – to maintain temperature at 25 °C 1.5 M barium chloride (BaCl2) – dissolve 366 g BaCl2 in d.i. H2O and dilute to 1 L – to precipitate absorbed carbonate in alkali solution as BaCO3 k. Phenolphthalein solution – dissolve 1.0 g phenolphthalein in 100 mL 95% ethanol – to serve as acid/base color indicator l. 1 M hydrochloric acid (HCl) – standardize against ~0.5 g (to nearest 0.0001) oven-dried TRIS [(HOCH2)3CNH2] using THAM indicator solution (0.1 g BromoCresol Green + 0.1 g Alizarin Red S in 100 mL water) – to neutralize alkali m. Digital dispensing burette – to dispense acid for neutralizing base n. Stir plate and stir bar – to mix alkali solution and keep BaCO3 away from acid

Method 1. Obtain representative soil sample from field of interest at defined depth (recommended depth of 0-10 cm, but other depths can be used if desired for specific purpose) 2. Sieve oven-dried (55 °C, 48-72 h) soil to pass a 4.75-mm screen (all material should pass the screen, except stones) 3. Weigh a 50 g portion of soil into a 60-mL graduated glass container. We often use two 50-g portions to obtain the flush of CO2 from 100 g soil. [This SOP describes an approach with standardized soil weight and variable volume, but an alternative approach to standardize volume and vary soil weight would be to use a 50 mL cup to which soil could be lightly packed and subsequently weighed to determine total porosity; both approaches require variable water delivery] 4. After all soil samples have been placed into glass containers, determine volume occupied by soil and lightly pack similar soils to the same volume, i.e. within nearest 2.5 mL. Add defined quantity of d.i. water to surface of soil and place glass container with soil into incubation jar. Amount of water should be 50% water-filled pore space (WFPS) – mL H2O = 0.50 x porosity x volume, where porosity is determined from weight and volume occupied by soil, e.g. 50 g soil occupying 40 mL volume has bulk density of 1.25 Mg m-3; resulting in 0.53 m3 m-3 porosity, therefore 40 mL volume requires 10.6 mL water to achieve 50% WFPS: Total porosity = (1 – BD/PD) where, BD = bulk density and PD = particle density (assume 2.65 Mg m-3)] 5. Add 10 mL of water to plastic vial and place in incubation jar [this is only to maintain humidity in jar] 6. Dispense 10 mL of 1 M NaOH into a labelled 30-mL Nalgene vial and place it along with glass container of soil into incubation jar [At this point the incubation jar contains a vial of water to maintain humidity, 1 or 2 glass containers of wetted soil to assess microbial activity, and a vial of alkali (NaOH solution) to capture evolved CO2] 7. Place completed box of 12 jars into a constant temperature room at 25 + 0.5 °C. Note: include one jar of an alkali blank without soil for every ~25 jars with soil, i.e. 5 blanks for every 100-sample set

8. At the end of 3 days of incubation, remove vial of alkali and immediately cap with air-tight screw-top cap 9. Once alkali vials from all incubation jars have been removed, open an alkali vial and titrate with HCl to a phenolphthalein endpoint (pink to colorless). First, add BaCl2 solution in excess (i.e., 1 to 3.3 mL) to alkali solution to form BaCO3 precipitate. Place a small magnetic stir bar into container and place vial on magnetic stir plate [stirring of solution is necessary to avoid reaction of HCl with BaCO3]. Add 3 drops of phenolphthalein indicator solution. Titrate with HCl until color changes from pink to clear/colorless. Record volume to nearest 0.01 mL 10. Calculate the quantity of C mineralized between 0 and 3 days from the equation: CMIN (mg kg-1 soil) = (mLBLANK – mLSAMPLE) x M HCl x 6 x 1000 / (g soil) where, 6 = equivalent weight of C and 1000 is conversion factor from g to kg Note: Each drop of acid (0.03 mL) is equivalent to 3.6 mg CO2-C kg-1 soil (50 g sample) or 1.8 mg CO2-C kg-1 soil (100 g sample), and therefore, this can be considered the minimum detection limit.

Costs o o o o o

Equipment (controlled temperature incubator - $5K, dispenser - $5K, digital burette - $1.2K, pipette - $300, 100 glass bottles, plastic bottles, Mason jars and lids, plastic vials - $400, stir plate - $100) – total = $12,000 Operating (NaOH, HCl, BaCl2, phenolphthalein for 100-sample batches) – total = $75 Space requirements = 5 m2 floor space and 8 m2 bench space, including a sink and ideally a hood for soil processing and analyses Time to get 100 CO2 values from start to finish = 5-6 days Labor to get 100 CO2 values = 26-30 hours

Sensitivity to management The flush of CO2 is responsive to differences in soil type [6, 7, 8], soil texture [2], tillage [5, 9, 12, 15], crop rotation [1, 15], fertilization [14], organic amendment [1, 19], land use [18, 19, 20], pasture management [13, 14, 15, 16], aggregate disruption [5], and soil sampling depth [12, 13, 15, 16].

Dynamic nature The flush of CO2 is somewhat seasonally dependent on root growth dynamics [1, 9], but more sensitive to yearly changes in organic inputs [13, 15, 16, 17] and highly sensitive to decadal changes in management-induced organic inputs [18].

Relationship to soil function The flush of CO2 reflects the functional capability of soil to cycle nutrients [4, 10, 11], decompose organic amendments, and catalyze and stabilize ecosystem processes through the interactions among a diversity of organisms. Soil microbial activity is solely from natural conditions without further amendments. Microbial activity from dried and coarselysieved soil is similar to that in dried and intact soil [2].

Production lab capability The method only requires basic laboratory equipment and the nature of the testing fits ideally to soil testing requirements for being rapid, inexpensive, reproducible, suitable for a wide range of soils, and correlating to nutrient needs of crops and meeting environmental goals [3, 4]. Variations of the concept are being used in different laboratories, including the Haney Soil Health Test

(https://www.wardlab.com/download/biotesting/Haney_Rev_v1_Information.pdf), Solvita Soil CO2 Burst Test (https://solvita.com/co2-burst/), and the Cornell Comprehensive Soil Health Assessment (http://www.css.cornell.edu/extension/soil-health/manual.pdf).

References [1] Culman SW, Snapp SS, Green JM, Gentry LE. 2013. Short- and long-term labile soil carbon and nitrogen dynamics reflect management and predict corn agronomic performance. Agronomy Journal 105, 493-502. [2] Franzluebbers AJ. 1999. Potential C and N mineralization and microbial biomass from intact and increasingly disturbed soils of varying texture. Soil Biology and Biochemistry 31, 1083-1090. [3] Franzluebbers AJ. 2016. Should soil testing services measure soil biological activity? Agricultural and Environmental Letters doi:10.2134/ael2015.011.0009. [4] Franzluebbers AJ. 2018. Soil-test biological activity with the flush of CO2: III. Corn yield responses to applied nitrogen. Soil Science Society of America Journal (accepted 15 March 2018; doi:10.2136/sssaj2018.01.0029). [5] Franzluebbers AJ, Arshad MA. 1997. Soil microbial biomass and mineralizable carbon of water-stable aggregates. Soil Science Society of America Journal 61, 1090-1097. [6] Franzluebbers AJ, Haney RL, Honeycutt CW, Arshad MA, Schomberg HH, Hons FM. 2001. Climatic influences on active fractions of soil organic matter. Soil Biology and Biochemistry 33, 1103-1111. [7] Franzluebbers AJ, Haney RL, Honeycutt CW, Schomberg HH, Hons FM. 2000. Flush of carbon dioxide following rewetting of dried soil relates to active organic pools. Soil Science Society of America Journal 64, 613-623. [8] Franzluebbers AJ, Haney RL, Hons FM, Zuberer DA. 1996. Determination of microbial biomass and nitrogen mineralization following rewetting of dried soil. Soil Science Society of America Journal 60, 1133-1139. [9] Franzluebbers AJ, Hons FM, Zuberer DA. 1995. Tillage and crop effects on seasonal soil carbon and nitrogen dynamics. Soil Science Society of America Journal 59, 1618-1624. [10] Franzluebbers AJ, Pershing MR. 2018. Soil-test biological activity with the flush of CO2: II. Greenhouse growth bioassay from soils in corn production. Soil Science Society of America Journal (accepted 15 March 2018; doi:10.2136/sssaj2018.01.0024). [11] Franzluebbers AJ, Pershing MR, Osmond D, Crozier C, Schroeder-Moreno M. 2018. Soil-test biological activity with the flush of CO2: I. C and N characteristics of soils in corn production. Soil Science Society of America Journal (accepted 15 March 2018; doi:10.2136/sssaj2017.12.0433). [12] Franzluebbers AJ, Schomberg HH, Endale DM. 2007. Surface-soil responses to paraplowing of long-term no-tillage cropland in the Southern Piedmont USA. Soil and Tillage Research 96, 303-315. [13] Franzluebbers AJ, Stuedemann JA. 2003. Bermudagrass management in the Southern Piedmont USA. III. Particulate and biologically active soil carbon. Soil Science Society of America Journal 67, 132-138. [14] Franzluebbers AJ, Stuedemann JA. 2005. Soil carbon and nitrogen pools in response to tall fescue endophyte infection, fertilization, and cultivar. Soil Science Society of America Journal 69, 396-403. [15] Franzluebbers AJ, Stuedemann JA. 2008. Early response of soil organic fractions to tillage and integrated crop–livestock production. Soil Science Society of America Journal 72, 613-625. [16] Franzluebbers AJ, Stuedemann JA. 2015. Does grazing of cover crops impact biologically active soil carbon and nitrogen fractions under inversion or no tillage management? Journal of Soil and Water Conservation 70, 365-373. [17] Haney RL, Hons FM, Sanderson MA, Franzluebbers AJ. 2001. A rapid procedure for estimating nitrogen mineralization in manured soil. Biology and Fertility of Soils 33, 100-104 [18] Jangid K, Williams MA, Franzluebbers AJ, Blair JM, Coleman DC, Whitman WB. 2010. Development of soil microbial communities during tallgrass prairie restoration. Soil Biology and Biochemistry 42, 302-312. [19] Jangid K, Williams MA, Franzluebbers AJ, Sanderlin JS, Reeves JH, Jenkins MB, Endale DM, Coleman DC, Whitman WB. 2008. Relative impacts of land-use, management intensity and fertilization upon soil microbial community structure in agricultural systems. Soil Biology and Biochemistry 40, 2843-2853. [20] Jangid K, Williams MA, Franzluebbers AJ, Schmidt TM, Coleman DC, Whitman WB. 2011. Land-use history has a stronger impact on soil microbial community composition than aboveground vegetation and soil properties. Soil Biology and Biochemistry 43, 21842193.