,Phosphorous (P) is one of the major macronutrient required for crop growth. It is important to know the amount of P that is available for plant uptake as P can limit crop productivity leading to low yield. Available P is very sensitive to pH therefore soil pH should be monitored so as not to limit P.
This video illustrates the method; check back because we are currently updating the method though the video will shows the main steps, which will not change:
You will need:
- Hanna high range field colorimeter (hannainst.com part number HI-717; note: we are using the low range reagents with the high range colorimeter. this is intentional).
- Reagent packet from Hanna : phosphate low range reagent (hannainst.com part #HI-93713)
- 25- mL graduated cylinder
- A small juice bottle, e.g. 100-500 mL, or a 50-mL centrifuge tube, for shaking soil with extractant, washed and rinsed with water free of phosphate (see below)
- A second small juice bottle for filtering (300-500 mL e.g.) with a wider-type mouth, about 4 cm diameter, with a modified lid that has holes punched in it with a medium sewing needle or a fine drill. You can see the complete filtration rig instructions, which may be useful for other filtering tasks, in the separate video below.
- Distilled water or phosphate-free water (this can be tested with the hanna kit and reagents, to verify that it has low phosphate levels, e.g. < 0.5 ppm phosphate; many bottled water brands that are not advertised as “mineral water” have these levels.
- Paper filters to filter the extract and produce a transparent soil extract for analysis, two alternatives.
- Coffee filters (filter paper, cone-shaped model) can be used on many soils that have lower clay content. They will initially pass clay particles, but after a few drops have passed the clay will plug the filter pores and clear solution will be obtained. This method can be slower but is accessible if laboratory-type filters cannot be found.
- Lab filters, qualitative #5 Whatman type, 90 mm diameter. This is the smallest-pore laboratory filter in the cheapest qualitative grade that can be found (#5, with an estimated pore size of 2.5 microns). If using another type of lab filter you should try it with typical soils from the region to check that it will not leave a cloudy solution (see the method below). Buying the 90-mm diameter will allow you to cut 4 smaller filter discs from one large filter disc, so that a box of 100 filters will work for 400 samples.
- Vials for reacting soil extract with the reagent packet (above) and reading the amount of blue color developed in the Hanna colorimeter: 11-mL vials, 3/4 “ in diameter to fit the colorimeter.
- Transfer pipet (or dropper) to transfer extract from the bottle or centrifuge tube to the filtering bottle after allowing to settle.
- Small plastic cups to receive filtered the extract and for acidifying extract (2 cups per test).
- A squirt bottle with P-free water to facilitate rinsing out cups and precisely adding water to the extract.
- Olsen solution: 0.5 NaHCO3 + enough NaOH to raise the pH to 8.5 (about a 2g NaOH per liter as a rough estimate)
- Solid NaHSO4 (sodium bisulfate) to acidify the Olsen soil extract in preparation for the color reaction.
- Measure 2.5 g soil into the 50-mL centrifuge tube or juice container. If you are using the #5 Whatman lab filters, the centrifuge tubes or a narrow bottle are best, to improve the settling out of clays.
- Using the 25-mL graduated cylinder (or a balance to weigh the solution), add 25 mL Olsen solution to the soil in the bottle. (solution can also be pre-weighed on the day of measurement)
- Shake 20 minutes
- Allow to settle for 10 minutes. For some soils this will produce a settling of most of the clay, for others only a little settling will occur. Still, this is useful to not have to filter so much clay which becomes time-consuming.
- Now you have a choice of filtering method depending on which type of filter you are using. the first option below is fastest and more foolproof if you have access to #5 whatman paper filters – especially for heavy clay soils– but the latter can be made to work with patience and a little practice, and uses only standard cone filters which are more available in many places.
Note: see the video below on the filtration rig as an idea for filtering the extracts, in addition to the filtering procedures below.
Filter method 1 with Whatman #5 lab paper filters:
- using a transfer pipet or dropper, carefully suck up the clearer liquid from the top of the tube or bottle and transfer this to the second bottle with a perforated lid (needle or drill holes). You should transfer off about 10 mL or a little more to allow for using 7 mL in the analysis below. Note that the settled extroct will not be completely clear, and also that if you do suck up a little of the unsettled clay it is alright. We only want to reduce the load of clay on the filter, not eliminate it.
- Place a filter of the right size in the lid that completely fills the lid area so that the edges will seal against the mouth of the bottle. This filter disc will probably need to be cut to fit, and you can make a thin cardboard pattern or even a cutting disc to make this faster and easier.
- Screw the lid with the filter onto the bottle and hand-tighten.
- Turn the bottle over and begin to squeeze. Drops of extract will begin to fall into the cup. With a #5 lab filter, these should be clear, so check for a torn filter if not.
- To avoid fatigue in squeezing, you can make a press like that shown in the video (YouTube video below), out of wood and long bolts, that suspends the bottle above the cup. This saves time because you can move to a new sample while leaving the first one in the press.
Filter method using cone coffee filters
- It is very helpful to practice the below method on an unimportant test sample to allow mistakes and learning to occur, prior to actually filtering a real sample. This method seems to work well for soils that do not lie at the clayey end of the soil textural triangle, i.e. with < 35% clay. For very clayey soils the #5 Whatman lab filter method above may be better, though either can be attempted.
- Starting from the 10 minutes of letting clay settle out in the Olsen solution, prepare the second bottle with a double layer of coffee filter in the lid with the holes. You can use a pattern or a circular cutter to mark and cut the filter dics. Cone filters come as a double layer, which facilitates just cutting the disc out of the doubled up cone.
- To transfer the extract to the filtering bottle In this method you can either use a transfer pipet to take 15 mL or so out of the extraction bottle which was shaken and transfer to the filtering bottle, or just pour directly and carefully from one to the other. In this method of filtering, you should actually be sure to transfer some of the clayey layer from the extract bottle to the filter bottle, because the filter needs to become a bit clogged with clay so that the clay stops passing through it to get a clear filtrate (again, trial and error will help here).
- Cap the filtering bottle with the perforated cap with the filter disc, and hand tighten.
- Turn over the filtering bottle and begin to squeeze, either by hand or using the filter press.
- Now it is important to watch the first 5 or 10 drops of filtrate that emerge. These drops should start out cloudy, and then will become clearer and clearer. The first cloudy drops are discarded. When the drops emerging are only transparent or a little yellow/brown and contain no cloudiness, you can switch to catching them for analysis. If the solution stays very muddy, it is likely the filter has ripped and you should reinstall another filter.
- Continue filtering until you have 7 mL of fluid, or more if desired. if filtering becomes very slow and you can see that the filter inside the bottle has developed a layer of clay, you can turn the bottle back on its side, remove the lid and the filter, and replace the clayey one with a clean filter. You will need to check again that the drops coming out of the filter turn clear before catching them for analysis.
- Now, regardless of the filtering method above you should have between 5 and 10 mL of sample, ideally 7 mL.
- Measure 7 mL (less if filtering was difficult) into the graduated cylinder from the cup that you used to catch the filtered extract (shake out any remaining drops out of the cylinder from measuring the clean Olsen solution that you mixed with the soil at the beginning)
- In a second dry cup, measure 0.45 g dry Sodium bisulfate. If this is a very calcareous soil, it is possible that more bisulfate will be needed (e.g. 0.50 g) since the extract will have a greater neutralizing potential (ie slightly higher pH because of the calcareous soil) for the bisulfate. Also, if less filtrate was used in step 9 above, you will need to reduce this amount, e.g. 0.32 g if only 5 mL was used versus 0.65 g for 10 mL filtrate (see table 2 below for exact amounts). These are targets, if you are within about 0.02 g of these amounts of sodium bisulfate, the color development should be relatively even among samples.
mL filtered …… Sodium bisulfate to add (g)
5 mL ……. 0.32 g
6 mL ……. 0.38 g
7 mL ……..0.45 g
8 mL ……..0.51 g
9 mL ……..0.58 g
10 mL ……0.65 g
- Add the filtered extract in the graduated cylinder to the cup with the bisulfate and allow to fizz and bubble as the bicarbonate reacts with the acid in the salt. We are bringing the extract pH from about 8.5 to below 5 or 6, so that when the reagents are added, they will bring the pH down further to about 1 or 2 where the blue color development of the Molybdate-Ascorbic Acid method can occur.
- While allowing the solution to fizz and gradually lose its bubbles, you can add the reagent package to the vial for the colorimeter, in step 13 below.
- Reintroduce the extract from the cup to the graduated cylinder. Rinse using the squirt bottle with a little water (<10 mL) and add this, eventually making up the volume to 20 mL in the cylinder. This will give you 10 mL for the reagent vial in the colorimeter, and 10 mL as a blank for the colorimeter (the extract will be a straw to brown color, so we want to correct for this color development in the colorimeter)
- Mix the contents of the graduated cylinder well by pouring back and forth between the cup from neutralizing step and the cylinder, two to three times. It can be surprisingly hard to mix liquids in a tall cylinder, so be sure to perform this step.
- Add the contents of the reagent package to a clean vial. You can cut the package straight across the very top, then open the top of the package to a square opening and pour along one corner of the square a few times, to avoid losing any reagent. This step gets better with practice.
- Add 10 mL of the solution to the vial with the reagents. You can create a vial as a pattern to copy by filling it with exactly 10 mL and marking the line to use as a guide for all the vials you use in analysis. Also, if you leave one vial cap’s thickness between the vial threads and the liquid meniscus, this is about 10 mL.
- Add the remainder of the solution in the cylinder of solution to a control vial. For samples from the same approximate soil type, a single control vial can be used for all the samples because the brown or straw color of these samples will all be similar. When large differences in organic matter occur, the control vial will be darker for higher organic matter soils and a different control vial should be used for this different set of samples.
- You may have to shake and uncap the reagent vial a few times to release bubbles. However, if adding the reagent causes immediate and vigorous bubbling, this may be an indication that insufficient sodium bisulfate was added in the previous step to fully neutralize the samples. If you know the approximate pH of the soil, you’ll find that this occurs for high-pH soils, and you can adjust the pH of the diluted sample in the cylinder with about 0.05 g more NaHSO4 (more is excessive) and remeasure.
- Blue color should develop in the reagent vial, while the control vial without reagent will retain its original color. Read the blue color after about 15-20 minutes on the Hanna colorimeter. Briefly, here are the steps for using the colorimeter:
a. Press button to turn on (press and hold to turn off first if just finishing another measurement)
b. Wait for C1, insert control vial, and press the button again.
c. Wait for C2, insert blue-colored vial with reagent, and press button again.
d. Take reading (see the active carbon protocol for more information on using the Hanna colorimeter).
- Log the reading. You may want to repeat the readings after 20, 25, and 30 minutes at first, to test whether the color development continues.
- In many soils, complexation and precipitation of the blue color with organic matter (OM) dissolved by the basic Olsen extraction will begin to occur after about 40 minutes, and sometimes sooner. Once this happens, the blue color will begin to drop, or may go up temporarily because the floating particles block the light path in addition to the blue color. It is important to “catch” the reading before lots of precipitation occurs and these blue flakes appear, and therefore 15 to 20 minutes is usually a good time.
- Disposal of the diluted reagents on an infertile soil will likely not cause any adverse effect or toxicity, and molybdenum may even act as a plant nutrient. In high salinity areas the sodium from sodium bicarbonate in the extracts could be a problem and it is best to dispose in a public sanitation system. If acidity is a concern, reagents can be neutralized back to neutral with a little wood ash or lime.
Calculation of final results:
here is how the concentration of available P is calculated (a version of Olsen P provided by soil labs).
- the concentration of “reactive phosphorus” in the solution of the colorimeter vial (not all of this is technically inorganic phosphate, since some organic P in solution also reacts with the reagents):
Raw concentration of P = Praw = 0.0559 x Reading – 0.0052
This is a calibration curve made by analyzing Olsen solutions with known concentrations of added phosphate. Then,
- Concentration of P in extract = [Pinorg] = Praw x (20 mL / N mL extract used in dilution),
where the “N mL extract used in the dilution” is the 7 mL added to the graduated cylinder (or N of between 5 and 10 mL as described in step 8-10 above) before acidifying and diluting. I.e., if we used the standard procedure above, we would multiply Praw by 20/7 or just 2.86 to get [Pinorg] This concentration has units of mg P / L.
- Amount Pinorg = mass of P in soil originally = [Pinorg] x 0.025 L (in mg P units)
(we multiply the concentration by the amount of solution, 25 mL or 0.025L, we extracted from)
Concentration of Olsen available P in soil , the result usually given in a soil test =
[Pinorg]soil = Pinorg / 0.0025 kg (units mg/kg) Or, we may have to adjust for the exact weight of soil we used at the start, e.g. 0.00243 kg if we used 2.43 grams dry soil.
Worked example for results:
Say we measured a result of 12.3 on the colorimeter, after extracting 2.52 g soil according to the procedure above, and using 7 mL of the filtered extract to then dilute up to 20 mL before measuring.
so, Raw P concentration measured by colorimeter = 12.3 x 0.0559-0.0052 = 0.6824 mg P/L
Concentration of P in extract before dilution = 0.6824 x 20/7 = 0.6824*2.86 = 1.952 mg P/L
Amount of P extracted = 1.952 x 0.025 L = 0.04879 mg
Concentration of P in soil = 0.04879 mg / 0.00252 kg = 19.4 mg P / kg
This would be an Olsen P result in the medium to high range.