Total Phosphorus in Plants and Soils: Digestion and Colorimetric Analysis

Edited by E. Herbert, December 2012

Last Updated by S. Widney, April 2016

  1. SAMPLE PREPARATION

Wear gloves for sample prep. Make sure sample material is oven dried or desiccated for 48 H before weighing.

  1. Print off TP analysis template (L-drive  Methods & Templates  Soil Phosphorus  Templates).
  1. Locate the metal test tube rack used for TP digestions ALWAYSput it in white plastic tub fill rack with 39 acid-washed, 100 mL tubes (2nd drawer down, below CHN analyzer) NOTE: Place the tube w/ thermometer in position 21 (see diagram). It is used as a separate check of digestion block temperature.

Front

  1. Put the tub of tubes next to the analytical balance  place a piece of weigh paper (on shelf above spec) on the balance  tare the weight weigh oven dried, ground and sieved (2 mm mesh diameter) sample.

For soils and woody plant tissue: 0.1995-0.2005g

For herbaceous/leaf tissue: 0.0995-0.1005g

  1. For each sample: Record tube #, sample id and weight on data sheetfold weigh paper together in a funnel shape transfer to corresponding digestion tube, leaving the paper in the top of the tube to mark your place. NOTE: If there is significant residual sample left on the weigh paper  place weigh paper back on balance  record weight & subtract from previous sample weight.
  1. You must also include a standard reference material (SRM) in tubes 37, 38 and 39 to determine the efficiency of the digestion. For

Forsoil: 0.1995-0.2005gof estuarine sediment

For plant tissue: 0.0995-0.1005g of pine leaves

  1. The last digestion tube will be used as a blank and is left empty until the digestion procedure. It will be treated like all the samples through the digestion and chemistry to account for any contamination.
  1. DIGESTION PROCEDURE

Protective gear required! Rubber gloves (or double-glove), lab coat, rubber apron, and face mask.

REAGENTS

  1. 500 mL bottle of concentrated nitric acid (HNO3;16N)repipetter
  2. 500 mL bottle of concentrated perchloric acid (HClO4; 70%)repipetter
  1. Place digestion block in the perchloric acid fume hood and place tube rack in the block. BE CAREFUL, it is easy to break the tubes when moving them in and out of the block.
  1. Check to make sure there is enough nitric acid in the bottle for all of the samples to be digested (total vol. = 5mL X # of samples, standards, and blanks). If necessary, fill the bottle with more acid before using.
  1. Check the accuracy of the repipetter by dispensing 3X into a 10 mL graduated cylinder, making sure it is consistently delivering 5 mL.
  1. Place rack of tubes in the digestion block. Pipette 5ml of Nitric acid into each digestion tube.
  1. Set block to 127C and record time when block reaches temperature. Check color, liquid level of samples and temperature (digital & thermometer) every 30 min.Record. (Remember to put on your protective gear when handling samples).
  1. Allow samples to digest for ~4 hours or until solution turns from red to yellow or clear. Record end time.

DO NOT ALLOW LIQUID LEVEL TO DROP BELOW SAMPLES.

NOTE:Organic soils could take as long as 16-20 hrs, but do not exceed 20 hrs. This can be done in multiple time blocks so that samples are not digesting overnight. When doing this, unplug digestion block. (probably will not take 16 hours, but could be longer than 4)

  1. Unplug the digestion block  allow the test tubes to coolcover with aluminum foil.
  1. Repeat steps #2-4 for the perchloric acid pipette 3 mL of HClO4 into each digestion tube.
  1. Place a glass funnel into each tube to enable acid to reflux. NOTE: it is easier if you can use all small funnels. If not, try alternating small and medium funnels.***THIS STEP IS EXPLOSIVE IF THE SAMPLES ARE COOKED TOO LONG. USE CAUTION AND CONSTANTLY MONITOR SAMPLES***
  1. Set digestion block temperature to 172C. Record start times when block reaches 172C allow samples to digest for 2 hours MAXIMUM (Should only take 1 hour). Check every 20 min. and record color, liquid level of samples and temperature (digital & thermometer). The samples should stop “poofing” when they’re done.

NOTE: Samples will “poof” steam and at the end of the digestion, the solution will be mostly clear and there may be some solid material in the bottom of the tubes.

  1. Unplug the digestion block  allow samples to cool for ~1 hour.
  1. Carefully remove rack from digestion block and place in the plastic tub and cover with foil. Allow to cool to room temp. (Easier to remove tubes one by one and set in extra rack rather than moving whole rack at once)
  1. Remove funnelsand rinse thoroughly in tap water IMMEDIATELY after removing them. It’s easiest to fill a plastic tub with water and transfer the funnels to the tub as you remove the tubes from the rack.
  1. Dilute each tube to mark (100 mL) with Milli-Q water. (Initially add water ~ halfway up in fume hood, then finish on benchtop).
  1. Cover each tube with a piece of Parafilm invert 3Xto make sure the sample is thoroughly mixed  repeat for all samples, standards, and blanks.
  1. Allow the tubes to sit over night so particulates will settle out. Put digestion block away.
  1. Make sure hood is closed completely and empty. Perform hood wash down.
  1. DIGESTATE STORAGE
  1. Label an acid-washed, 50 mL polycarbonate centrifuge for each sample digested (with marker, no tape). Label the foam tray(s) with a piece of tape indicating: Sample type, TP digest, Name, Date
  1. Pour a 50 ml aliquot of sample into its respectivecentrifuge tube store samples at 4° C until analysis.
  1. Dispose of left over digests in labeled TP waste container  immediately rinse digestion tubes with tap water 3x and scrub thoroughly with the scrub brushNOTE: Tubes may need to be scrubbed with some Neutrad water to remove soil stuck to the tube.
  1. COLOROMETRIC ANALYSIS

REAGENTS

NOTE: Check for all reagents, glassware, consumables and cuvettes several days prior to analysis. Plan for colorimetry to take ~6 hours (including cleanup)

All reagents should be at room temperature before using in analysis.

  1. Phenolphthalein Indicator: weigh out 0.05 g phenolphthalein  transfer to a 50 mL volumetric flask  dissolve and dilute to mark with 50 ml EtOH and 50 ml Milli-Q water cap & invert several times to mix  store in a dropper bottle in “toxic” cabinet beneath hood. Expires in 2 months. Please note your name, date made, and date of expiration on solution. (Okay to use after expiration)
  1. 5N NaOH: Weigh out 50 g of solid NaOH pellets into a plastic weigh boat. Transfer the pellets to a labeled 250 mL volumetric flask (helps to use a wide funnel). Add a stir bar to the flask and put it in an ice bath on a stir plate in the fume hood. Slowly add water to the flask while stirring (hold on to the flask so it doesn’t tip over in ice bath) until the flask is mostly full (just below the neck). Allow the solution to stir until all of the pellets are dissolved and the solution is not hot to the touch (~10 minutes). Remove stir bar and dilute to mark with Milli-Q water cap & invert several times to mix  store in a plastic bottle in “base” cabinet beneath hood. Does not expire. Please note your name and date made on solution.
  1. 1000, 100, and 10 mg P/L Standards:
  2. Dry ~ 1 g of KH2PO4 at 105°C for 1 hour
  3. Weigh and quantitatively transfer 0.439 g of dried KH2PO4 to a 100mL volumetric flask  add ~ 80 mL of Milli-Q water & dissolve completely.
  4. Dilute to mark with Milli-Q water  cap & invert several times to mix thoroughly  store in a glass bottle in the fridge. Expires in 6 months. Please note your name, date made, and date of expiration on solution.
  5. Make 100 ppm KH2PO4 solution from 1000 ppm solution by pipetting 10 ml of 1000 mg P/L standardinto a 100 ml volumetric flask, diluting to the line, and inverting to mix. Expires in 6 months. Please note your name, date made, and date of expiration on solution.
  6. Make 10 ppm KH2PO4 solution from 100 ppm solution by pipetting10 ml of 100 mg P/L standardinto a 100 ml volumetric flask, diluting to the line, and inverting to mix. Expires in 6 months. Please note your name, date made, and date of expiration on solution.

NOTE: All standards should be stored long term in the fridge at 4°C, but allowed to warm to room temperature before using

  1. Solution A, Ammonium Molybdate/Potassium Tartrate: *CAUTION*This solution should be made in the hood WEARING PROTECTIVE GEAR & the volumetric flask should be put in a cold water bath b/c this reaction is extremelyexothermic!Use ice packs from the freezer and a plastic tub for the ice bath.
  2. Add ~ 400 mL of Milli-Q water to a 1 L flask  weigh and transfer 50g of ammonium molybdate and 1.213g of antimony potassium tartrate to the flask.
  1. Measure 500ml of concentrated sulfuric acid (H2SO4), using a graduated cylinder. Slowly (25-50 mL at a time), transfer measured acid to flask. Wait until solution has completely cooled before adding more.
  1. When all the acid has been added,dilute to mark with Milli-Q water, cap & invert several times to mix thoroughly.Store in a 1 L glass Pyrex bottle in the fridge @ 4° C. Expires in 6 months. Please note your name, date made, and date of expiration on solution.
  1. Solution B, Ascorbic Acid:
  2. Weigh and transfer 8.8g of ascorbic acid to a 100 mL volumetric flask  dilute to mark with Milli-Q water  cap and invert to mix thoroughly. NOTE: If ascorbic acid solution turns yellow, discard and make new. If crystals in reagent bottle turn yellow, discard and purchase new.Made fresh day of assay. Wrap volumetric flask in aluminum foil to protect from light.
  1. Working Solution/Color Reagent:

*CAUTION* This solution should be made in the hood & fresh for each analysis

  1. Add ~ 500 mL of Milli-Q water to a 1 L volumetricflask measure & quantitatively transfer 100 mL of Solution A (ammonium molybdate, antimony potassium tartrate) and 100 mL ofSolution B (ascorbic acid) to flask  dilute to mark with Milli-Q water  cap & invert several times to mix thoroughly.Made fresh day of assay.
  2. Cover flask in foil and add stir bar. Stir on low setting until solution is used.
  1. ANALYSIS – CALIBRATION CURVE
  1. Take the cover off the Shimadzu spectrophotometer  turn it on by flipping the switch on the rear, left side  let warm up for ~ 30 minutes.
  1. Make sure the wavelength says “882 nm”. If not press the “go to WL” button  press clear 3 X to completely erase the current wavelength  type “882” using the # pad  press “enter” to set the wavelength  the screen should now display 882 nm above the “abs” reading.
  1. Rinse a quartz cuvette3x in Milli Q water.
  1. Make calibration standards (make fresh for every analysis):
  1. Determine the concentration range that will bracket the concentration of P in your samples  select ~ 5-6 standards to make within that range  label 50 mL volumetric flasks accordingly.
  1. Below is a suggested set of standards and how to make them. Remember to calibrate the pipette using Milli Q water each time you change the volume.

Concentration of standard Volume of 10 mg P/L Std. Added to 50 mL Flask

0 mg P/L 0 mL

0.2 mg P/L 1 mL

0.4 mg P/L 2 mL

0.6mg P/L 3 mL

0.8 mg P/L 4 mL

1.0 mg P/L 5 mL

1.2mg P/L6 mL

  1. Add the specified volume of 10 mg/L standard to its respective flask. Add a small amount of water (10-20 mL) to each flask. Add two drops of phenolphthalein to each flask, then add 5N NaOH until pink color appears (should not take much NaOH).
  2. Add 20 mL of working solution to each flask, fill to the line with Milli-Q water, and invert 3 times to mix.
  3. Allow standard to react for 30 minutes before measuring absorbance (*see details below). NOTE: Standards should turn increasingly blue with increasing concentration.
  1. Zero the spec with Milli-Q water. Analyze standards, from lowest to highest concentration. Invert 1st standard a few times to mix rinse quartz cuvette with standard 1x, and then fill ¾ full, handling it from the toptap the cuvette on the counter gently to remove bubbles → wipe down the sides with a Kimwipe place cuvette, w/ clear window facing the light path (frosted side facing you), into the spec position listed on the screen (right-hand side)  close the door once the value has stabilized, record the absorbance on your data sheet.
  1. Pour standard solution in cuvette into waste container  rinse cuvette with ultrapure water and leave it out for reading samples.
  1. Repeat steps #1 & 2 for all calibration standards.
  1. Go back to the “TP analysis template” file  enter the absorbance readings for the standards in the 2nd worksheet in the file  generate a calibration curve with regression equation & r2 check equation with previous TP runs to make sure the slope and intercept are comparable & the r2 must be at least 0.99 or better before proceeding with your samples.
  1. ANALYSIS - SAMPLES
  1. Label a 50 mL volumetric flask for each sample to be analyzed.
  1. Using a 10-mL pipette, pipette either 10 mL (samples high in TP; e.g., plants) or 20 mL (samples low in TP; e.g., soils) of sample into respective flask  record volume on data sheet.
  1. Add 2 drops of phenolphthalein indicator to each flask.
  1. Neutralize samples as follows:
  1. On 1stsample, use an eyedropper and slowly add small amounts (uL) of 5 N NaOH and swirl constantly. Keep track of the volume and stop once the solution turns barely pink (i.e., endpoint). NOTE: You do not want to overshoot the endpoint b/c it will screw up the colorimetric chemistry.
  1. Now you can neutralize subsequent sample a little faster by adding a bulk of the volume added to the 1st sample, then adding one drop at a time until you reach the endpoint.
  1. Using a 10-mL pipette, add 20 ml of working solution to each flask  dilute to mark with Milli-Q water  cap & invert several times to mix thoroughly.
  1. Allow samples to react for 30 minutes before measuring absorbance. Only dose ~13 samples at a time, and stagger the sets of samples by ~30 minutes for best timing.
  1. Measure & record the absorbance for your samples by following the same procedure used for the calibration standards.
  1. Enter the data for your samples on the same datasheet as your calibration standards  save the file w/ an appropriate name on the L-drive
  1. The remaining solution in the flasks should be poured into a labeled TP waste container (in room w/ hoods)  flasks should be rinsed with tap water  put into “dirty glassware” to be acid-washed.
  1. DATA ANALYSIS
  1. Minimum R2 of 0.9990 needed for standard curve.
  2. To calculate TP concentration (mg P/L) in the 50 mL volumetric flask from the absorbance reading you record on the spec, you must use the regression equation generated from your calibration curve. NOTE: It is an easier calculation if you graph absorbance on the x-axis and concentration on the y-axis.
  1. Y (conc in 50 mL flask.; mg P/L) = slope*absorbance + intercept
  1. To calculate TP concentration (mg P/L) in the digestion tube from the concentration in the 50 mL volumetric flask (step #1), you must account for the dilutionfactor:
  1. TP (digestion tube; mg P/L) = TP (flask; mg P/L) * (50 mL/xmL)
  1. x mL = volume of sample pipetted into 50 mL volumetric (e.g., 10 or 20 mL; make sure to use consistent units)
  1. To calculate ug P/g sample, take the concentration determined in step #2 and do the following calculation:

------

Sample weight (g)

  1. To calculate % recovery for the standard reference materials (SRMs), use the following equation:
  1. SRM Measured TP (mg/L) = SRM TP (step #2; mg/L)
  1. SRM Expected TP (mg/L) = (SRM (g) * (%P/100)) * 1000 mg/g

------

0.100 L

  1. % recovery = Measured TP (mg/L)

------* 100

Expected TP (mg/L)

Alternative to above: Determine “Fudge Factors” to plug in to data sheet:

Sample type / Sample weight / x / Dilution in digestion tube / x / Dilution in volumetric flask / = / Fudge Factor
Soil / 1 g/(~0.2 g) / x / 100 mL / x / 50 mL/20 mL / = / 1250
Soil / 1 g/(~0.2 g) / x / 100 mL / x / 50 mL/10 mL / = / 2500
Plants / 1 g/(~0.1 g) / x / 100 mL / x / 50 mL/10 mL / = / 5000

*Bold denotes the most commonly used fudge factors.

To generate values of ug P/g, multiply the measured concentration (ppm) by the appropriate fudge factor.

Calculating Percent Recovery using the NIST Standards

Three NIST standards are to be processed and measured as a part of each Total Phosphorus run performed. These are stored in the desiccator. Examine the type of sample you have, look in the NIST standard book (on the book shelf) to verify that the NIST you would like to use has a certified measurement of phosphorus. The Phosphorus content is often written on a % mass fraction basis.

To determine your percent recovery take the three values of P (ug/g) measured on your NIST standards (often these are Sample #s 37-39).

Following from our above example:

These samples are plant samples, as such, the “Trace Elements in Pine Needles”

standard was used.

The NIST standard gave a Mass Fraction (%) of 0.107 +/- 0.008 for Phosphorus.

The colorimetric procedure above measured the following three values in units of

(ug P /g soil) for the NIST standard:

1053.108, 1002.092, 987.516

Following simple stoichiometry convert these to Mass Fraction (%)by dividing by 10,000. This should result in the following three values:

0.1053108, 0.1002092, 0.0987516

Divide each of these by the NIST standard (0.107) to yield the following respective (%) recoveries:

98.42%, 93.65%, 92.29%

Digestion procedure is adapted from:

Sommers, L.E. and D.W. Nelson. 1972. Determination of total phosphorus in soils: a

rapidperchloric acid digestion procedure. Soil Science Society of America Proceedings. 36: 902-904.

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