Soluble Protein Assay to Quantify Soluble Protein Content in Plant Tissues

To quantify soluble protein content in a plant tissue, in a tube, begin with an appropriate amount of ground, lyophilized plant tissue.

Add sodium hydroxide, an alkali, to the tube and sonicate. The alkaline environment with the mechanical forces disrupt the cells, releasing intracellular contents, including macromolecules - soluble proteins and carbohydrates. Incubate at higher temperatures.

In the presence of heat, alkali cause protein hydrolysis, forming smaller peptides and increasing protein solubility. Additionally, carbohydrates get degraded.

Centrifuge the mixture. Collect the solubilized protein-containing supernatant. Add hydrochloric acid to neutralize the pH.

Treat the sample with trichloroacetic acid, TCA. TCA disrupts the hydration shell surrounding the proteins, leading to protein aggregation and precipitation.

Centrifuge and remove the TCA-containing supernatant. Wash the protein pellet with ice-cold acetone to remove any residual TCA, which may interfere with soluble protein estimation.

Air-dry the acetone. Resuspend the protein pellet in sodium hydroxide. Dilute with deionized water. Transfer to the wells of a multi-well plate.

Add an acidic solution of Coomassie Brilliant Blue G-250 dye. Under acidic conditions, the dye binds to the basic amino acid residues in the solubilized proteins, resulting in a blue protein-dye complex.

Use a spectrophotometer to measure the absorbance of the protein-dye complex, proportional to the soluble protein content in the plant tissue.

To begin, weigh out replicate samples of each tissue into labeled 1.5-milliliter microcentrifuge tubes. Next, record the precise mass of each sample. Using a micro-pipettor, add 500 microliters of 0.1 molar sodium hydroxide to each tube. Close the lids tightly, and sonicate the tubes for 30 minutes.

Next, place the tubes in a pre-heated hot water bath for 15 minutes. After this, centrifuge the tubes at 15,000 g for 10 minutes. Pipette the supernatant into new labeled microcentrifuge tubes using a new pipette tip for each sample.

Add 300 microliters of 0.1 molar sodium hydroxide to the pellet, and repeat the centrifugation for 10 minutes. After this, remove the supernatant, and transfer it to the tubes containing the supernatant from the first centrifugation.

To neutralize the pH of the supernatant, add 11 microliters of 5.8 molar hydrochloric acid. Use litmus paper to confirm the pH is approximately 7.

Next, add 90 microliters of 100% trichloroacetic acid to each tube. Then, incubate the tubes on ice for 30 minutes. Centrifuge the samples at 13,000 g for 10 minutes at 4 degrees Celsius. Carefully, use a vacuum line and glass micropipette tip to remove the trichloroacetic acid.

Be very careful not to disturb the pellet with the suction tip.

Next, wash the pellet with 100 microliters of acetone cooled to negative 20 degrees Celsius. Then, allow the acetone to evaporate in a fume hood. Dissolve the protein pellet with 1 milliliter of sodium hydroxide. Additional rounds of heating, vortexing, and sonicating may be required.

Add 160 microliters of each IgG standard solution to a 96-well plate in triplicate, starting with the A1 position. Next, in a new 1.5-milliliter tube, add 50 microliters of each sample solution to 950 microliters of distilled water. Then, add 60 microliters of each diluted sample to the well plate in triplicate, starting with the H1 position. Add 100 microliters of distilled water to all of the blank and unknown sample wells.

Using a multi-channel pipette, add 40 microliters of Coomassie brilliant blue G-250 protein dye to each well of the plate. Using a needle, pop any bubbles present in the wells. After this, allow the plate to incubate at room temperature for 5 minutes. Then, use a microplate spectrophotometer to record the absorbance values for each well at 595 nanometers.