Documentation:Soil Permanganate Oxidizable Carbon
What is Active Carbon?
Active carbon is a labile pool of soil organic carbon that typically represents 3-4% of total soil C, has been identified as a good indicator of soil quality (Cambardella and Elliot, 1994). This pool of SOC is easily decomposed and is associated with microbial activity, nutrient cycling, and formation of soil aggregates. It is also sensitive to changes in land management, and can be used as a sensitive indicator of soil quality (WWeil et al., 2003)
How can we measure Active Carbon?
One relatively inexpensive and less environmentally damaging method to determine active carbon in the soil is a method developed by Weil et al. (2003) using potassium permanganate. The following protocol has been developed through the use of that protocol and the protocol established by Culman et al. (2012).
Permanganate Oxidizable Carbon
- Distilled water
- 70% ethanol (likely made from 95%)
- Reagent grade potassium permanganate
- 0.1 M NaOH
- Soil samples ground and sieved <2mm
- 100-1000ul volume micropipette and disposable tips
- 50ml centrifuge tubes with lids (2 per sample, 13 for standards)
- 2 - 500ml beakers
- 1000ml beaker
- 2 - funnel
- 2 - 1000ml volumetric flasks
- 2 - stir bars
- Stir plate / hot plate
- pH meter
- Spectrophotometer (needs to be able to read 550nm absorbance
- Obtaining and Prepping Samples
- Samples should be obtained from the field at the 0-7.5cm area. Typically three soil samples will be compiled mixed, then subsampled to obtain a good representation of the field. (If you are taking aggregate stability samples, it may be reasonable to use a portion of sample that you obtain for this analysis).
- Store your samples in the fridge until you are able to process them.
- Air dry your samples, grind, and sieve to <2mm.
- Preparation of materials
- Ensure that all materials have been washed using acid bath prior to set up.
- Ensure all materials that come in contact with KMnO4 solution are washed after use, as residue can accumulate on them.
- Label 2 falcon tubes for each sample (sample name a, sample name b)
- Label 9 falcon tubes with the concentrations used for the standard curves:
- 0.005 (a, b), 0.010 (a, b), 0.015 (a, b), 0.020 (a, b), and a blank for water.
- Label 4 falcon tubes for the standard samples
- Soil standard (a, b) and solution standard (a, b)
- Stock KMnO4 preparation (makes 1 L of 0.2 M KMnO4)
- Appropriate safety gear including gloves, lab coat, and safety glasses should be worn when using KMnO4)
- Each soil sample analyzed will use approximately 2.0mL of the 0.2 M KMnO4 solution.
- Weigh 147 g of CaCl2 into a 1000mL volumetric flask and dissolve in deionized water using a stir bar and stir plate. Once fully dissolved, bring solution volume up to 1000mL.
- Weigh 31.60g of KMnO4 powder into a 1000mL beaker and add approximately 900mL of the CaCl2 solution. Using a magnetic stir bar and a stir plate on a gentle heat (~3 setting or between Low and Medium), stir until KMnO4 is completely dissolved. Note: The dissolution may be very slow, and due to the very dark colour of the solution it may be necessary to decant some of the solution to check for undissolved KMnO4.
- Using the calibrated pH meter, adjust the pH of the KMnO4 solution by adding 0.1 N NaOH one drop at a time. Since only a few drops will be needed, allow the pH reading to stabilize between additions of NaOH (the endpoint approaches rapidly!). Once the pH has been adjusted, pour the solution into a 1000mL volumetric flask and bring the volume to 1000mL with the remaining CaCl2 solution. Note: Try to avoid spilling any CaCl2 solution, as almost all of it will be needed to bring the KMnO4 solution up to volume. pH 7.2
- Transfer the finished KMnO4 solution to an amber bottle (since the solution is photosensitive) and store in a dark cupboard. The solution can last from 3-6 months.
- Extraction procedure
- Add 49.5mL distilled water to 50mL centrifuge tubes labeled 1b-10b, Soil Std (b), Solution Std (b), and the ‘b’ standard curves (including water blank). Cap all of the tubes and set aside.
- Place a labeled centrifuge tube (‘a’ tubes) onto scale and tare. Measure out 2.50g (+/- 0.02g) of soil sample into the tube and record mass of soil. Cap the tube after filling.
- Using the dispensette, add 18mL of distilled water to the soil sample tubes (‘a’ tubes), as well as the tubes labeled Soil Std (a) and Solution Std (a).
- Pour approximately 60mL of 0.2 M KMnO4 solution into a 500mL beaker, and return solution to the cupboard.
- **Time is now of the essence in the procedure**
- Note the time. Using the 1000µL micropipette, pipette 1.0mL (1000µL) of 0.2 M KMnO4 into tubes 1-10, in increasing order. Add a second 1.0mL of 0.2 M KMnO4 in decreasing order, from 10 to 1. Cap the tubes tightly, hand shaking each for 1-2 seconds and bring to the shaker. Note: Store the beaker with the KMnO4 solution in a dark cupboard or turn off the lights when leaving the room.
- Note the time. Secure the tray with tubes 1a-10a horizontally on the shaker (see figure below) and shake on the ‘High’ setting for 2 minutes. After 2 minutes, turn off the shaker, remove the tubes and transport them back to the workbench.
- Note the time. Invert each tube once to clean soil off the sides of the tubes, then uncap the tube and set gently back in the tray. Once all the tubes are uncapped, place the tray in a dark cupboard and allow the suspension to settle from 10 minutes from the time that was noted at the start of this step.
- Just before the 10 minutes is up, uncap the water-filled tubes labelled 1b-10b.
- After 10 minutes, gently remove the incubating sample tubes from the dark cupboard.
- Remove 0.5mL (500µL) of the supernatant from tube 1a and add it to the distilled water in tube 1b. Repeat for the rest of the samples in increasing order, using a new micropipette tip for each sample. Note the time when you finish extracting from the last tube.
- Cap the tubes tightly and invert gently to mix. Set tubes aside a dark cupboard until ready to fill well plate for the spectrophotometer (Section 3.5). Note: If extracted properly, the solutions should be stable for up to 12 hours if kept in the dark as they should no longer be reacting with soil carbon (Culman et al., 2012).
- Note: In successive extractions, try to keep the time intervals (between time to fill, shaking, incubating, extracting, etc.) as consistent as possible to minimize time sensitivity errors.
- Stock solution and working standards
- Stock Solutions
- Pre-wet a 10mL glass pipette with distilled water.
- Add 9.0mL distilled water to the remaining 50mL centrifuge tubes labelled 0.02, 0.015, 0.10, and 0.005, using the 10mL pipette.
- Using the 100-1000µL micropipette, add 750, 500, and 250µL distilled water to the tubes labelled 0.005, 0.010, and 0.015, respectively.
- Tightly cap the tubes and invert their content to mix. Store in the dark unless you are preparing the working stands immediately after this step.
- Working Standards
- Gather the tubes labelled 0.02, 0.015, 0.10, and 0.005, that were initially filled with 49.5mL of distilled water in Section 3.3.a.
- Using the large micropipette, add 0.5mL (500µL) of the 0.005 M KMnO4 stock standard to the tube containing distilled water to make the working standard.
- Continue with each stock standard, making one working standard for each KMnO4 concentration (see figure below).
- Stock Solutions
- Reading on the spectrophotometer
- Pipette 250µL triplicates from each soil sample centrifuge tube (‘b’ tubes), from the working standard solutions, the Soil Std (b) tube, and the Solution Std (b) tube into a 96-well microplate.
- When finished, cover the microplate with a lid and read on the TECAN spectrophotometer in MCML 240. If necessary, the plate can be stored in a dark place for up to 12 hours (Culman et al., 2012).
- Turn on the TECAN spectrophotometer and warm up for 2-3 minutes.
- Turn on the computer attached to the machine, and logon (ask TA or instructor for credentials)
- Open the TECAN SPARKCONTROL application, found on the desktop, and run the “Soil Active C” method. Follow prompts to load the plate (lid should be off). When finished, copy data to a USB stick and save to a folder on the computer.
- Refer to Culman et al. (2012) for setting up a spreadsheet for absorbance analysis.
- A plate map should be prepared ahead of time designating which sample or standard has been placed into which well.
- Before pipetting into the microplate, invert the centrifuge tube 2-3 times to ensure proper mixing of the solution.
- For access to MCML 240 and the TECAN spectrophotometer, e-mail the FNH lab coordinator (Imelda Cheung, firstname.lastname@example.org)
- At the end of section 3.3.k, the solutions are dilute enough to be disposed of by pouring down the sink and flushing with generous amounts of water.
- Dispose of KMnO4 contents to a bottle labelled “Waste KMnO4” and cap the bottle tightly. This can be mixed with waste soil and left in a bright area until it becomes a light pink colour. The solution can then be safely disposed of down the drain followed by generous amounts of water.
- 50mL centrifuge tubes should be soaked in an acid bath, rinsed with distilled water and air-dried before being used for the analysis. After completion of the POXC analysis, tubes should be rinsed thoroughly with tap water as soon as possible and soaked overnight in the acid bath. If left sitting with KMnO4 solution, a residue can accumulate on the inside of the tubes.
- Culman, S., Freeman, M. & Snapp, S. (2012). Procedure for the Determination of Permanganate Oxidizable Carbon. Retrieved 16 December 2016 from https:// s3.amazonaws.com/metadata_production/protocols/133/KBS+POXC+Protocol.pdf
- Weil, R., Islam, K., Stine, M., Gruver, J., & Samson-Liebig, S. (2003). Estimating active carbon for soil quality assessment: a simplified method for laboratory and field use. American Journal of Alternative Agriculture, 18(1), 3-17.