13.11: Lab Technique - SDS-PAGE

Separating Proteins by Denaturing SDS-PAGE Electrophoresis

Electrophoresis (meaning to “carry with electricity”) separates molecules based on their size using an electric field. Like DNA, electrophoresis can separate proteins based on their charge and size. While agarose can be used in this separation, the preferred polymer is polyacrylamide. Polyacrylamide is a water-soluble polymer made up of linear chains of acrylamide. It can be cross-linked and polymerized to form a soft gel comprised of small pores capable of separating proteins ranging from 5 to 250 kDa. The separation of proteins using polyacrylamide is known as Polyacrylamide Gel Electrophoresis or PAGE. Because most PAGE techniques use a detergent, such as sodium dodecyl sulfate (SDS), to denature the proteins being separated, the separation of proteins by electrophoresis is often called SDS-PAGE or denaturing SDS-PAGE. PAGE in the absence of detergents is called native-PAGE. SDS-PAGE separates proteins primarily by their size because the ionic detergent SDS denatures proteins into their individual subunits and binds to each subunit to make them uniformly negatively charged. Thus, when a current is applied, all SDS-bound proteins in a sample will migrate through the gel toward the positively charged electrode. The smaller protein subunits will migrate faster through the gel, while the larger proteins will take longer to migrate towards the positive electrode. Therefore, the rate at which a protein subunit passes through the gel is inversely proportional to its size.

Preparation of the Polyacrylamide Gel

Polyacrylamide gels are prepared by mixing acrylamide with bis-acrylamide in a defined ratio. The amount of polyacrylamide in the mixture produces gels of specific percentages (i.e., concentrations). The higher percentage of acrylamide in the gel translates to smaller the pores within the gel and better resolution of small proteins (5 to 60 kDa). For example, a 7% acrylamide gel will have larger pores in comparison to a 10% gel. Low percentage gels are used to separate larger proteins (e.g., from 60 to 200 kDa). The standard polyacrylamide gel used is a 10% gel which can separate proteins ranging from 20 to 150 KDa. However, "gradient" gels can be prepared with higher percentages of acrylamide (i.e., 20% with smaller pores) at the bottom of the gel and lower percentages (i.e., 4% with larger pores) at the top. This enables the user to separate a larger range of proteins using a single gel. The polyacrylamide-bis-acrylamide mixture is crosslinked to form a gel through the addition of a polymerizing agent, ammonium persulfate (APS) and the catalyst TEMED (N,N,N’,N'-tetramethylenediamine). The gels are often prepared in a buffer that is similar to the one that will be used to run the gel. Because of this, this type of SDS-PAGE is called discontinuous.

Once prepared, the acrylamide/bis-acrylamide solution is poured between into a thin space found between two glass or plastic plates. These plates are assembled with spacers of specific size to form a gel with a desired thickness. The assembled plates are called a "cassette" and the process is referred to as "casting" a gel (Figure \(\PageIndex{1}\)). To obtain optimal resolution of proteins, two types of gels are made and polymerized in the cassette: a separating or resolving gel with a specific concentration and a stacking gel with a set concentration of 7%. The separating gel makes up the majority of the gel's total volume (i.e., 80%) and separates the proteins according to their size. The stacking gel has a lower pH and a different ionic concentration than the separating gel. This, together with the discontinuous nature of the system, ensures that all proteins, irregardless of their size, flow through the stacking gel at the same rate and enter into the separating gel at the same time. A stacking gel is not necessary when using a gradient gel, as the gradient itself performs this function. The separation gel is poured into the cassette first and allowed to polymerize. The stacking gel is then poured on top of the resolving gel. A "comb" is inserted into the stacking gel and allowed to polymerize. When the comb is removed, wells are created for individual protein samples.

Users now have the option of buying "pre-cast" gels for SDS-PAGE. Precast gels are a more convenient way of analyzing proteins and give researchers a level of gel consistency that hand cast gels may not have. Precast gels are available in a range of percentages and include difficult-to-pour gradient gels. Precast gels are also available in the different buffer formulations (e.g., Tris-glycine, Bis-Tris, Tris-acetate), which are designed to optimize shelf life, run time, and/or protein resolution.

Preparing the Samples

In denaturing SDS-PAGE, samples are prepared in a concentrated Sample Buffer containing the anionic detergent SDS and a reducing agent. The presence of SDS denatures proteins into their subunits and surrounds them with a uniform negative charge. Reducing agents, like dithiothreitol (DTT) or β-mercaptoethanol, break disulfide bonds found between cysteine amino acids, denaturing the protein subunits into their linear conformations. Denaturing is assisted by heating the protein sample 70-100°C. Finally, typical Sample Buffers contain some type of tracking dye (usually bromophenol blue) used to assess the rate of sample migration through the separating gel.

Running the Gel

Once the gels polymerize, it is ready to be "run" (Figure \(\PageIndex{1}\)). To run the gel, the cassette is mounted (usually vertically) into an electrophoresis tank containing an upper cathode and a lower anode. Once the cassette is placed in the tank, an upper and lower buffer chamber will form and the top and bottom edges of the gel cassette will come into contact with the cathode and anode, respectively. The comb is removed from the stacking gel, the wells rinsed, and the two buffer chambers in the electrophoresis tank are filled with a Running Buffer. The most common Running Buffer is a combination of Tris base and Glycine. The Tris-Glycine buffer allows for the efficient flow of electricity through the gel and permits the separation of a large range of protein sizes. Alternate buffers include Tris-Acetate, Tris-Tricine, or Bis-Tris. The Tris-Tricine buffer is a modification of the Tris-Glycine buffer. It is often used with gels that resolve low molecular weight proteins in the range of 2–20 kDa. Tris-Acetate and Bis-Tris buffers, with their neutral pH, decrease protein degradation in the gel and are often used in the analysis of high molecular weight proteins or native-PAGE applications where the more neutral pH is beneficial. However, gels run with Bis-Tris buffer must be run with alternative reducing agents, such as sodium bisulfite. The samples are loaded alongside a "protein ladder" of known molecular weights in order to assess the size of the separated proteins. The gel is run at a constant voltage (e.g. 120V to 150V) until the tracking dye found in the Sample Buffer is reaches off the bottom of the plate.

Figure \(\PageIndex{1}\): Denaturing SDS PAGE workflow. (1) The separating gel is poured between two glass plates and allowed to polymerize. Isopropanol is layered on top of the separating gel to create a smooth interface between separating and stacking gel. (2) The isopropanol is removed and the stacking gel is poured. A comb is inserted to create wells. The gel is allowed to polymerize. (3) The comb is gently removed and the wells rinsed with PBS or water. (4) The prepared electrophoresis "cassette" is loaded into the electrophoresis tank and the chambers of the tank are filled with Running buffer. The protein samples are loaded into the wells. (5) The gel is run. (6) The cassette is removed from the tank. The gel is removed from the cassette and can be stained to visualize protein bands. If moving on to a Western blot, the gel is not stained. (SDS PAGE by Patricia Zuk by CC BY 4.0; figure created in BioRender. Zuk, P. (2025))

The protocol given below outlines gel electrophoresis using precast or hand-cast denaturing SDS-PAGE gels.

Lab Protocol: Gel Preparation

1. For a precast gel:
   1. remove the gel from its packaging, remove the comb and rinse wells with deionized water.
2. For a hand cast gel:
   1. Assemble the plates to form the gel cassette.
   2. Prepare a separating gel and pour into the cassette. The volume of this gel should be approximately 80% of the total volume of the cassette.
   3. Add a thin layer of isopropanol to prevent evaporation and create a smooth interface with the stacking gel.
   4. Allow the gel to fully polymerize.
   5. Pour off the isopropanol and rinse with deionized water.
   6. Prepare the stacking gel and pour into the cassette until the volume reaches the top of the notched plate.
   7. Insert the desired comb size.
   8. Allow the gel to fully polymerize.
   9. Gently remove the comb and rinse the wells with deionized water.

Lab Protocol: Sample Preparation

1. Prepare the proteins samples for analysis by mixing with a 4X Sample Buffer (i.e. Laemelli Buffer) in a 3:1 ratio.
2. Boil the samples for 95°C for 5 minutes.
3. Allow to cool.

Lab Protocol: Electrophoresis

1. Secure the gel cassette in the electrophoresis tank with the notched plate facing inward.
2. Fill the upper and lower buffer chambers with 1X SDS Running Buffer.
3. Using protein loading tips, carefully pipette the samples into the appropriate wells. Ensure one of the wells is a protein ladder for reference.
4. Connect the electrophoresis tank to the power supply.
5. Run the gel at 120V until the tracking dye reaches the bottom of the gel (~1-2 hours).
6. Remove the cassette from the tank.
7. Carefully separate the two plates of the cassette.
8. Scrape off the stacking gel. and discard.
9. For visualization of protein bands, stain the separation gel. Do not stain if Western blotting. Refer to Western Blotting protocol for additional steps.

Lab Protocol: Gel Staining and Visualization

1. Carefully remove the separating gel from the cassette plate and immerse in Coomassie Brilliant Blue staining solution.
2. Incubate for 30 minutes with gentle agitation.
3. Destain the gel by washing in Destaining solution with gentle agitation. Several changes of this solution may be necessary. (e.g., methanol/acetic acid/water mixture). Destain until protein bands are visible and the background is clear.
4. Compare the migration of protein bands to the molecular weight marker to estimate protein sizes.

Safety Considerations

• Handle acrylamide with care as it is a neurotoxin in its liquid form. Use gloves and work in a well-ventilated area.
• Dispose of buffers and gels according to institutional waste disposal guidelines.

PAGE Reagents

30% Acrylamide/0.8% bis-acrylamide stock

• add 150 g of acrylamide to 250 mL distilled water and mix
• add 4.0 g bis-acrylamide and mix
• add distilled water to 500 mL and mix well
• filter through a 0.2 µm filter
• store at 4ºC in a foil-wrapped bottle

10% Separation gel (pH 8.7)

• combine the following:
  • 5.0 mL 30% acrylamide/0.8% bis-acrylamide solution (10%/0.3% final)
  • 3.75 mL 1.5M Tris-HCl, pH 8.7 (0.375M final)
  • 150 µL 10% SDS
  • 75 µL 10% APS
  • 7.5 µL TEMED
• add distilled water to 15 mL
• pour gel immediately

4% Stacking gel (pH 6.8)

• combine the following:
  • 2.0 mL 30% acrylamide/0.8% bis-acrylamide solution (4%/0.1% final)
  • 3.75 mL 0.5M Tris-HCl, pH 6.8 (0.125M final)
  • 150 µL 10% SDS
  • 75 µL 10% APS
  • 15 µL TEMED
• add water to 15 mL
• pour gel and insert comb immediately

5X SDS Running Buffer (pH 8.3)

• combine the following in 800 mL distilled water:
  • 15.1 g Tris base
  • 72.0 g glycine
  • 5.0 g SDS
• add water to 1 L (pH will be ~8.3)
• store at room temperature
• dilute to 1X before use
• 1X final concentration:
  • 25 mM Tris
  • 208 mM glycine
  • 0.1% SDS

4X Sample Buffer

• combine the following:
  • 250 µL 0.5M Tris-Cl pH 6.8
  • 400 µL 10% SDS
  • 250 µL 80% glycerol
  • 100 µL 1M DTT or 25 µL β-mercaptoethanol
  • 50 µL 1% bromophenol blue
• store at room temperature
• 1X final concentration:
  • 31.25 mM Tris-Cl pH 6.8
  • 1% SDS
  • 5% glycerol
  • 30 mM DTT or 0.005% β-mercaptoethanol
  • 0.0125% bromophenol blue

Coomassie Brilliant Blue Staining Solution

• dissolve 1.0 g Coomassie Brilliant Blue R250 in 300 mL methanol
• add:
  • 50 mL glacial acetic acid
  • 650 mL distilled water
• filter through Whatman paper
• store at room temperature protected from light
• final concentration:
  • 0.1% Coomassie Brilliant Blue R250
  • 30% methanol
  • 5% acetic acid

Destaining Solution

• to 650 mL of distilled water add:
  • 300 mL of methanol
  • 50 mL of glacial acetic acid
• store at room temperature
• final concentration:
  • 30% methanol
  • 5% acetic acid