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Unit 4
Analysis of Proteins by Electrophoresis
Introduction:
Electrophoresis is the process of placing proteins in a matrix and applying an electric field. This will separate the proteins according to their size, shape and charge. Gels made from polymerized acrylamide—polyacrylamide gels—are frequently used because they have high resolving power, can use relatively larger amounts of protein, do not interact with the proteins and have a stable matrix. Typically the polyacrylamide is crosslinked with N,N’-methylene-bis acrylamide in a polymerization reaction that is initiated by the molecule N,N,N’,N’-tetramethylethenediamine (TEMED). If low concentrations of acrylamide and bis-acrylamide are used, the pores are larger allowing for analysis of higher molecular weight proteins. Conversely, smaller molecular weight molecules are analyzed in gels with higher concentration of acrylamide and bis-acrylamide. The entire complex forms a matrix of fibers as shown in Fig. 1 below, where represent the polyacrylamide and the represent the crosslinking:
Fig. 1: Matrix of Polyacrylamide with Crosslinking
The PAGE gels are polymerized between two square panes of glass that are sealed around the edges and stood upright during polymerization. The resulting gels are run upright as well, giving rise to the common name of “vertical gel electrophoresis”.
PAGE gels may be continuous or discontinuous. Continuous gels have a gradient of acrylamide and bis-acrylamide concentration with the most concentrated gel at the bottom of the gel.
More often, a discontinuous gel is used. In this gel the lower 80 percent of the gel is called the resolving gel. This gel usually contains a higher percentage of crosslinks and is poured first. On top, there is a stacking gel that contains fewer crosslinks and prepared with a buffer containing fewer mobile ions. A comb is inserted when pouring the stacking gel to produce the wells into which the samples are placed. The stacking gel has lower resistance than the resolving gel so proteins will travel quickly through the stacking gel and concentrate at the interface. This allows the proteins to enter the resolving gel close together and therefore increases the resolution.
Native gels
When an electrical field is applied to the proteins they will travel down the gel. Their movement is influenced by many factors: size, shape, and charge. Smaller proteins will move more quickly through the gel than larger ones due to less frictional drag. For the same reason, tightly coiled globular proteins will travel more quickly than more loosely packed proteins or extended structure fibrous proteins in an electrophoretic gel. The ratio of charged amino acids in a protein will also influence movement through the gel; those proteins with more negatively-charge (acidic) amino acids than positively-charged (basic) amino acids will travel more quickly towards to positive anode due to their large net negative charge. A protein with a net positive charge will, in fact, migrate in the wrong direction from the sample well, away from the anode. Native gels when the conformation of the protein needs to be preserved, usually because the location of the protein of interest must be identified by its structural function. For example, if an enzyme must be identified by its enzymatic activity, a native gel will be used. Also, Western blots are typically performed with native gels to ensure that the antibody detection of a protein of interest will work.
Denaturing gels (SDS-PAGE)
With all these factors influencing the movement of native proteins through the PAGE (polyacrylamide gel electrophoresis) gel, it is nearly impossible to predict where a given protein will migrate or to analyze the results of a native gel. In order for proteins to behave similarly, they must be denatured into a uniformly extended shape. That way the unfolded proteins will behave similarly to nucleic acids in that the short polypeptides travel further down the gel and the longer polypeptides do not travel as far. Unlike DNA molecules, however, a denatured protein does not have a uniformly-charged structure. In order for denatured proteins to move similarly to each other in an electrophoretic gel, they must have a uniform charge-to-mass ratio.
There are two protein denaturing agents that are commonly used together in PAGE electrophoresis: a sulfhydryl reducing agent and an ionic detergent. Ditihiothreitol and 2-mercaptoethanol are short molecules containing a sulfhydryl (-SH) group that will break up the disulfide bridges and help to desrupt any tertiary structure and/or quaternary structure of the proteins due to disulfide bridges. (The sulfhydryl group is what gives these compounds their “rotten egg” smell.)
The main denaturing agent is an ionic detergent, sodium dodecyl sulfate or SDS. This detergent is the sodium salt of a 12-carbon alkyl sulfate compound. SDS actually serves two functions. First it disrupts the secondary, tertiary, and quaternary structures of the protein by breaking the ionic and hydrogen interactions between the amino acids of the protein, as well as interfering with the hydrophobic interactions responsible for correct folding of the protein. Secondly, SDS will coat the denatured proteins’ hydrophobic amino acid side groups with its hydrophobic docedyl tail, thereby coating the protein with negative charges from the sulfate head group of the detergent.
Typically, 1% SDS and 0.1 M mercaptoethanol are used along with high temperatures to completely denature proteins and coat their extended structures with negative charges. This gives rise to the SDS-PAGE electrophoresis that is the most common method of identifying an unknown protein or determining its molecular weight. In order to aid with this, a set of proteins markers of known molecular weight are commonly run along with the unknown. Analysis of sizes of protein bands in the gel is then a straightforward comparison to the migration distances relative to the molecular weight markers.
The samples of protein are diluted with a “sample buffer.” In addition to the the mercaptoethanol and SDS denaturants, the sample buffer contains a tracking dye that will travel with the front and determine how far the gel has run. Glycerol is also included for increased density of the sample in order for it to settle into the bottom of the well and not go floating off into the electrophoresis buffer. Typically somewhere between 10 and 40 mg protein are loaded into a well, depending on the purity of proteins in the sample. This total volume is typically about 20-40 mL, half of which is protein and half is the 2X sample buffer.
Lab 4-A:
Polyacrylamide Gel Electrophoresis
Introduction:
In the beginning of this unit there was a brief discussion of polyacrylamide gel electrophoresis and the different components used. Review this before coming to class.
Tonight we will be running a native gel and a SDS-PAGE gel on the purified green fluorescent protein (GFP) samples from Unit s 2 and 3. We will load two identical gels with identical samples, but samples with be prepared and run under two conditions: denaturing and nondenaturing. Once the gels are run, we will examine both under a UV lamp to look for fluorescence by the GFP band. We will then stain all of the protein bands with Coomassie blue, a stain for proteins. The Coomassie blue will stain the gel as well as the proteins, so the finishing step is destaining to remove the Coomassie blue that did not react with the protein bands.
Safety:
1. The wires connecting the cell to the power supply must be in good condition, not worn or cracked. Broken or worn wires not only cause rapid changes in resistance that adversely affects electrophoresis, but they also create an electrocution hazard.
2. Make sure the area around the power supply is dry.3. The area for at least 6 inches around the power supply and cell should be bare of clutter and other equipment. Clear space means any fire or accident can be more easily controlled.
4. Wear gloves while loading and handling the gels; the unpolymerized acrylamide is a neurotoxin.
5. Coomassie blue will stain clothing and hands. In addition it is very acidic. Wear gloves when handling the staining and destaining solutions. Lab coats or aprons are recommended.
Protocol:
Part I: Preparing solutions
Students may collaborate in the preparation of the following solutions for use by the entire class. Part of this collaboration is the sharing of the Media Prep Form describing the preparation of each solution in for individual lab reports. Each prepared solution should be labeled with its contents and concentrations, as well as a control number that cross references the solution to its Media Prep Form. Students using a solution should record its control number in their lab report.
Part II: Preparing the gels
1. With gloved hands, carefully un-wrap the cellophane from two PAGE gels, which are enclosed in a gel support chamber. The gel support chamber consists of thin plastic plates sandwiching the gel. One gel will be run under non-denaturing conditions, while the other will be run under denaturing conditions.
2. Handle the support chamber gently to avoid distorting the gel within. Drain any excess water out of the wells (the indentations in the top of the gel), and then clamp the gel support chamber onto the electrophoresis apparatus according to the instructions provided.
3. Pour the appropriate running buffer (see table below) into the top half of the apparatus to check for leaks. If sufficiently sealed, no buffer will appear in the bottom chamber. If this is the case, continue to fill the top of the apparatus until the gel and electrodes are covered with buffer.
Component / Concentration in the SDS-PAGE running buffer (1X) / Concentration in the native gel PAGE running buffer (1X)Trizma base / 25 mM / 25 mM
Glycine / 192 mM / 192 mM
SDS / 0.1% / -----
NOTE: The final pH of the running buffers should be 8.3.
4. Gently remove the comb from the stacking gel.
5. Rinse any unpolymerized acrylamide from the wells by pipetting the running buffer in and out of each well. The wells are too thin to use micropiptter tips; you will need to use special gel loading tips. Take care not to puncture the sides or the bottom of the well with your pipetter tip. This takes a steady hand – it may help to support the micropipettor with your other hand and to support your elbows on the lab bench top.
Part III: Preparing and loading samples into the gel wells
1. Defrost your samples of GFP from Labs 2 and 3.
2. Label microfuge tubes and add 20 mL of each GFP sample with an equal volume of 2X sample buffer. Choose GFP samples to represent different stages of your purification process. If purified GFP eluted from your column in multiple fractions, select the fractions that had the greatest amounts of GFP fluorescence. The components of the 2X sample buffer are given below. You need 11 x 20mL of each for two 10 well gels.
Component / Concentration in the 2X SDS-PAGE sample buffer / Concentration in the 2X nateive gel PAGE sample bufferTris-Cl pH 6.8 / 126 mM / 126 mM
Glycerol / 20% / 20%
Bromophenol blue / 0.0025% / 0.0025%
SDS / 4% / -----
3. Pipet up and down to ensure mixing. If your protein MW standards are not in sample buffer, add 10 uL to an equal volume of 2X sample buffer as well.
4. Incubate SDS-PAGE samples and protein MW standards in a 60° C water bath for 5 minutes. Dry the tubes with a paper towel and place them in the microcentrifuge in balanced positions. Centrifuge for 5 sec to force the solutions to the bottoms (simply turning the centrifuge on and immediately turning it off is sufficient.)
5. The wells are identified as #1 through #10, starting with #1 on your left as you face the front of the gel.
6. Read the Good Laboratory Practice Tips (below) and load 30 mL of each sample into a separate well and record the contents of each loaded well. Be sure to use a new gel loading tip for each sample loaded. Load each of the two gels in the same order.
GLP tips:To load a sample into a well,
a. Adjust an automatic pipetter to deliver the correct amount of the sample
b. Attach an ultrathin gel loading tip.
c. Withdraw the correct amount of your sample from your microcentrifuge tube and insert the pipetter tip into the top of the well to at least four mm of the bottom of the well. Take care not to puncture the sides or the bottom of the well with your pipetter tip. This takes a steady hand – it may help to support the micropipettor with your other hand and to support your elbows on the lab bench top. Make sure that the pipette tip is between the glass sandwich of the gel and very slowly and gently expel the solution from the pipetter tip into the well while holding the pipetter steady. The blue solution should fall to the bottom of the well, gradually filling it.
d. Do not press the pipetter to the second stop – it is important to avoid blowing air bubbles into the well.
e. Do not release your thumb until you have slowly withdrawn the pipetter tip from the well, so that you avoid removing the sample that you have so carefully loaded!
Troubleshooting:
¨ If the sample overflows into the adjacent well, you may be trying to load too much sample. Alternatively, you may be expelling the sample with too much force, or not withdrawing the micropipettor tip enough to make room for your sample as it fills the well. You may quickly withdraw any sample that has overflowed into an empty well.
¨ Work quickly to minimize the diffusion of samples from the wells or between wells.
7. Place a new tip on the automatic pipetter, then follow the procedure just described to load your “MW standards” into one well of each gel.
Part IV. Running your gels
Check to make sure that the upper buffer is not leaking into the lower chamber. If it is, you must disassemble the apparatus and fix the leak with vacuum grease along the rubber sealing ring. If the upper buffer chamber is not leaking, add running buffer to the lower chamber until the bottom of the gel is submerged.