The experimental design is a randomized block, with 5 replicates per treatment.  Within each block were 3 plots measuring approximately 1m x 1m, with a central circular area of 85 cm diameter (approx.  0.57 m2) used for treatments and sampling.  The treatments were as follows:

 

(C) control plots (no manipulations), 

(W) Nanopure water added to field capacity to 10 cm depth (5.6 L/plot), 

(WS) sucrose added in solution (15.82 g sucrose in 5.6 L water per plot).

 

Soil samples were taken for chlorophyll-a analysis as follows:  Sampling bags were prepared with one plastic vial and clean plastic teaspoon per sample.  Samples were taken from within the 85 cm diameter circular area of each plot starting at the 12:00 position on the first day.  The above treatments were then applied to the plots.  The 3:00, 6:00 and 9:00 positions were then sampled for each successive date. Soil was collected to 1 cm depth with the plastic spoon, excluding rocks with a diameter >5 mm.  About 4 teaspoons of soil were placed into the vial, keeping the vial out of the light in a closed hand.  The soil samples were stored in a light-tight bag, in a cooler for transportation.  On return to the laboratory (within 8 hours of sampling), the soils were stored at +5 degrees C until further processing.

 

Extraction of chlorophyll from the soil.  All procedures were carried out in the dark or very low irradiance to avoid degradation of the chlorophyll.  The soil samples were mixed thoroughly in the vials, and a sample of approximately 5 g was weighed out into a 50 mL plastic centrifuge tube with a screw-top cap.  10 mL of a 50:50 DMSO/90% acetone solution was added to each sample and they were mixed thoroughly on a bench-top Vortex mixer for about 5 seconds.  The vials were placed in a -4 degrees C constant temperature room, in the dark, and left for 12-18 hours.  

 

Determination of chlorophyll-a concentration.  This was determined fluorometrically using a Turner model 111 fluorometer.  A calibration using a known concentration of chlorophyll was carried out prior to sample analysis.  The machine was blanked using a 50:50 DMSO/90% acetone solution.  Each vial was mixed thoroughly, then centrifuged for 5 minutes at about 1800 RPM.  A sample of approximately 4 mL of the DMSO/acetone solution was taken from the top of the sample with a pipette, being careful not to get any soil particles in the solution.  The sample was placed in a cuvette, into the fluorometer and the fluorescence was recorded.  This was done fairly quickly in order to prevent light from breaking down the chlorophyll.  This measurement is called Fo, the initial fluorescence.  After taking this reading, 0.1 mL of 1N HCl was added directly to the cuvette and the cuvette was gently agitated.  After 20 seconds, the fluorescence was re-measured.  (During this step, the acid converts the chlorophyll to phaeophytin by releasing a magnesium ion in an acidic environment). This measurement is called Fa, the fluorescence after acidification.  The solution was discarded in to a waste container, and the cuvette rinsed 3 times with DMSO/90% acetone solution before proceeding with the next sample. 

 

Calculation of chlorophyll concentration.  The calibration curve that was constructed for the fluorometer had the following equation:  

 

 

The Fo and Fa figures were put into this equation to calculate chlorophyll concentration.  Subsequently, this figure was divided by 1000 to convert to microg/mL. Next, this was multiplied by 10 as the soil was extracted in 10mL of DMSO/90% acetone. Finally, this was divided by the fresh weight of the soil in g to give the concentration in microg chlorophyll per g fresh weight of soil.  If the extract was diluted prior to reading on the fluorometer, the dilution factor (noted in the comments column) was applied at the end of the equation.