Team:Cambridge/Protocols/Gel Electrophoresis of Protein

Protein Analysis by SDS PAGE

These are some notes made in preparation for running SDS PAGE to verify we have Reflectin.

Like DNA gels PAGE gels can be made with various different weight percentage SDS for different resolutions.

A 12% PAGE gel which we used will separate 12kDa-60kDa proteins.

Theory

SDS-PAGE or sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis is a method of resolving proteins of different molecular weights (kDa) by mixing samples with SDS, loading samples into usually an acrylamide gel and passing an electric current through it.

SDS is an anionic detergent which denatures secondary and non–disulfide–linked tertiary structures, and applies a negative charge to each protein in proportion to its mass, allowing fractionation of proteins via electrophoresis similar to a DNA gel with longer proteins experiencing more difficulty moving through the gel than shorter proteins. Samples are often heated in boiling water prior to loading to shake up the molecules and allow improved binding with SDS. A tracking dye for example bromophenol blue is used to indicate the stopping point.

After electrophoresis the gel is rinsed in D.I water and stained with a dye commonly coomassie blue, PAGEBlue or silver staining for improving fainter bands for visualisation of the separated proteins. After staining the gel is rinsed again and left to de-stain to your desired amount either in D.I water or in a de-staining solution.

There are protocols to make gels of the desired percentage weight of SDS and there are also pre-cast gels available from Bio-Rad and Life Technologies. We ran our gels using pre-cast 12% gels sourced from Bio-Rad using Bio-Rad electrophoresis tanks.

Practice

Before You Begin

Various buffers must be prepared prior to running a gel. If you are making your own gels you will need to make up all the buffers. If you are running a pre-cast gel then you will only require the 4x Sample Buffer, Reservoir Buffer and 4x Upper Tris Buffer. Please note that protocols for making gels vary but they all use the same ingredients.

In addition you will need a protein marker.

Reservoir Buffer

750 mls of 4x Tris-Glycine reservoir buffer-SDS

30 mls of 10% SDS

dH₂O to 3l

(Stable indefinitely at room temperature)

4x Tris-Glycine Reservoir Buffer - SDS

36g Tris Base

172.8g glycine

dH₂O to 3l

(Stable indefinitely at room temperature)

4x Upper Tris Buffer (0.5M Tris-HCl, pH 6.8, 0.4% SDS)

To 110 mls dH₂O add 12.12 g of Tris Base

Add 8 mls of 10% SDS

Adjust pH to 8.8 with HCl

Add dH₂O to a final volume of 200ml

(This solution can be stored for months in the refrigerator)

4x Lower Tris Buffer (1.5M Tris-HCl, pH 8.8, 0.4% SDS)

To 110 mls dH₂0 add 12.12 g of Tris Base

Add 8 mls of 10% SDS

Adjust pH to 6.8 with HCl

Add dH₂O to a final volume of 200ml

(This solution can be stored for months in the refrigerator)

10% Ammonium Persulfate Solution (TEMED)

0.1g Ammonium Persulfate

1ml dH₂O

(Stable for up to a month in the refrigerator)

4x Sample Buffer

4 mls glycerol

2 mls 2-mercaptoethanol

1.2g SDS

5mls 4x Upper Tris

0.03g Bromophenol Blue

(Aliquot into 1.5ml microcentrifuge tubes. Store at -20^oC. One tube can be stored at 4^oC for up to a month)

The SDS-PAGE Process

• Gel Making

PAGE gels comprise two parts; a lower resolving gel and an upper stacking gel. The stacking gel is where you load your samples into and in general is twice as tall as the desired wells. The lower resolving gel is where the protein bands will run through and be separated.

In addition the buffers above you will also require a casting assembly.

Resolving or Separating Gel

1. Clean a set of glass plates for each gel first with dH₂O and then with 70% ethanol
2. Assemble the casting assembly. Be sure that the bottom of the gel plates and the spacers are perfectly aligned. Place a mark on the glass plates 1.5cm below where the top of the gel will be.
3. Have ready:
   • a Pasteur pipette with a bulb
   • a P-1000 set to 1ml and a tip
   • ethanol
4. Prepare the separating gel solution according to bipartite tables 1 and 2.

Table 1: Preparation of the separating gel solution for 2 gels using 0.75 mm spacers

	5%	7.5%	10%	15%
dH2O	5.8 mls	5 mls	4.2 mls	2.5 mls
30% acrylamid/0.8% methylene bisacrylamide	1.7 mls	2.5 mls	3.3 mls	5.0 mls
4x Lower Tris	2.5 mls	2.5 mls	2.5 mls	2.5 mls
10% Temed	100 μl	100 μl	100 μl	100 μl

Table 2: Resolution ranges for gels of different acrylamide concentrations

Acrylamide Percentage	Separating Resolution
5%	60 - 212 kDa
7.5%	30 - 120 kDa
10%	18 - 75 kDa
15%	15 - 45 kDa

Gently mix the separating gel solution. Excess aeration will interfere with the polymerization of the gel.

dd 10μl of Temed to the separating gel solution. Mix well. Use paster pipette to fill the glass plates with separating gel solution up to the mark on the glass plates. Work rapidly as soon as the Temed is added because the gel will begin to polymerize upon addition of the Temed.

Over lay the gel with 1ml of the 95% ethanol

Let the separating gel polymerize for 30-60 minutes.

Stacking Gel

1. While the separating gel is polymerizing, prepare the stacking gel according to Table 3.

Table 3: Preparation of stacking gel solution for 2 gels using 0.75mm spacers:

Component	Volume
dH2O	3.25 mls
30% acrylamide/0.8% methylene bisacrylamide	0.5 mls
4x Upper Tris	1.25 mls
10% Temed	50 μl

Gently mix the stacking gel solution. Excess Aeration will interfere with the polymerization of the gel.

Get ready a Pasteur pipette with a bulb

Pour the ethanol off the separating gel. Rinse the gel assembly out with several changes of dH₂O.

Add 5 μl of Temed to the stacking gel solution. Mix well. Use a Pasteur pipette to fill the glass plates up to the top with stacking gel solution.

Carefully insert the comb. Be sure that no air bubbles are trapped to the ends of the teeth.

Let the stacking gel polymerize for 30 minutes.

There is an excellent youtube [http://www.youtube.com/watch?v=EDi_n_0NiF4&feature=related tutorial] which walks you step-by-step how to make the gel once you have made the solutions.

• Preparation of Protein Samples

SDS-PAGE can be run with pure protein samples, mixed protein samples and also directly from cell lysates. The general method for preparing proteins prior to loading is as follows:

1. Check the volume of the wells in the gel. Mix protein samples in volume ratio of 3:1 with 4x Sample Buffer in a microcentrifuge tube including the protein marker.
   • Note: If you don't have enough samples to completely fill the number of wells dilute the 4x Sample Buffer with D.I water and load this.
2. Submerge the protein in boiling water bath (100^oc works for me) for 2 mins.
3. Centrifuge for 20 secs at 12,000 rpm. You are now ready to load your samples into the gel.

• Running the Gel

1. (Load Gels) - Insert gels into electrophoresis tank making sure you have good contact with the smaller glass plate inwards.
2. (Load Reservoir Buffer) - Fill the inside and outside chambers of the electrophoresis tank with reservoir buffer. Fill the inside chamber up to brim.
3. (Remove Well Comb) - Having been lubricated by the buffer, carefully remove the combs forming the wells.
4. (Load Protein Samples) - Carefully load protein samples using a protein loading tip or pasteur pipette taking care there are no bubbles in the wells and samples are not flowing into other wells.
5. (Run Gel) - Connect the tank with the recommended power supply for your tank and set it to run at a constant voltage of 200V. Stop when the bromophenol blue is at the bottom of the tank.

Note: A fantastic [http://www.youtube.com/watch?v=XUjLO-ek2C8 video protocol] showing the process is available if you are unsure.

• Staining the Gel

There are a wide variety of protocols for staining the gels for visualisation after electrophoresis which vary depending on the stain used. Here we present a protocol for using Bio-Rad Bio-Safe Coomassie Blue G250 for staining. When staining place gel in a container that is approximately the same size as it.

1. (Remove the Gel) - Remove gel carefully from the tank and wash in D.I water (this will help later to remove the gel). Carefully pry the two glass slides apart taking care not to rip the gel. The gel will be stuck onto one of the slides. Gently submerge the slide with the gel still attached in D.I water. The gel will naturally fall off the slide. Remove the remaining slide leaving only the gel in the water.
2. (Washing the Gel) - Wash gel by incubating in D.I water bath for 5 minutes on a rocking table at a gentle speed. Repeat this step 3 times with fresh water. This step removes remaining SDS.
3. (Staining) - Submerge the gel with 50 ml Coomassie Blue G250 for 1 hour on a rocking table at a gentle speed for minigels. For larger gels use sufficient volume as to cover the gel. The resulting gel should be stained in blue and have well-stained protein bands.
4. (Destaining) - Submerge the gel in a D.I water bath on a rocking table at gentle speed overnight. A neat trick for faster de-staining is to introduce tissue balls in the bath but not directly on top of the gel to absorb the stain.

If you have faint bands you may also want to do a silver stain. Note: A fantastic [http://www.youtube.com/watch?v=b-1dXzU4iOw video protocol] is available if you are unsure. BEWARE: In the video they use PAGEBlue not coomassie blue.

Safety

The safety implication of the procedure.