Two-dimensional polyacrylamide gel electrophoresis (2-D PAGE) is a widely used technique for analyzing the proteome of yeast cells. To prepare yeast samples for 2-D PAGE, several steps are required to extract and solubilize the proteins.
The first step is cell lysis, which can be achieved by mechanical disruption using glass beads or by enzymatic digestion using lytic enzymes such as zymolyase. The choice of lysis method depends on the type of yeast and the properties of the proteins to be extracted.
Once the cells are lysed, the protein extract can be further purified and solubilized using different techniques, such as ammonium sulfate precipitation, phenol extraction, or organic solvent extraction. These methods can remove contaminants and improve protein solubility, which is important for 2-D PAGE.
After the protein extract is purified and solubilized, it can be separated using 2-D PAGE, which involves two steps of electrophoresis. In the first dimension, proteins are separated based on their isoelectric point using isoelectric focusing (IEF). In the second dimension, proteins are separated based on their molecular weight using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). The separated proteins can then be visualized by staining with a protein dye, such as Coomassie Blue or silver stain.
To identify the proteins on the gel, different methods can be used, such as peptide mass fingerprinting or tandem mass spectrometry. These methods can provide information on the identity, quantity, and post-translational modifications of the proteins.
Yeast samples for 2D PAGE can be prepared in (1) an initial cell disruption step; (2) a protein solubilization step; (3) a nuclease treatment step; and (4) a final stage in which the protein extract is diluted in IPG rehydration buffer before being applied to the first dimension of the 2D PAGE.
The procedure is carried out in 1.5-mL microcentrifuge tubes. A suitable starting material is a pellet of yeast cells from 10 mL of culture with a density of 5–10 million cells/mL, corresponding to an optical density (at 610 nm) of approx 0.5, which will typically yield a pellet of 5–10 μL of cells. The method described below is adjusted to this amount of cells.
The protocol can be scaled up or down; however, care should be taken to not use a final extract volume of more than 500 μL or less than 50 μL, since this will reduce the efficiency of the cell-disruption step.
1. Cell Disruption
A. Add 160 μL of ice-cold milliQ-quality water containing protease inhibitors to the cell pellet.
B. Add 0.25 g of chilled glass beads (diameter 0.5 mm) to the sample..
C. Vortex 4 × 30 s on a table shaker at maximum speed (approx 2500 rpm) with intermittent placement of samples on ice for at least 1 min.
2. Protein Solubilization
A. Add 20 μL of sample buffer I and vortex the tube(s) briefly to mix.
B. Place tube(s) at 95°C for 5 min. Make sure to secure the lid of the tube, alternatively to
C. make a small hole in the lid, prior to the heating step.
D. Cool samples on ice for 5 min.
3. Nuclease Treatment
A. Add 20 μL of sample buffer II and vortex the tube(s) briefly to mix.
B. Incubate on ice for 10 min.
C. Centrifuge samples at full speed (approx 15,000g) in a microcentrifuge at 4°C.
D. Aspire the supernatant (constituting the protein extract) to a new microcentrifuge tube and freeze at –20°C, or use immediately.
4. Dilution in IPG-Rehydration Buffer
A. Dissolve sample in required volume of IPG-rehydration buffer.
B. Incubate the sample at 37°C for 10 min.
C. Spin down sample (15,000g, 10 min, room temperature) to remove any particles that might remain.
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Reference
- Walker, J. M. (Ed.). (2005). The proteomics protocols handbook. Humana press.
