Analysis of Proteins by Western Blots

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Unit 3

Analysis of Proteins by Western Blots

Introduction:

Electrophoresis

Electrophoresis is the process of a biological sample into a molecular matrix and applying an electric field, providing excellent resolution of complex mixtures commonly used as an analytical technique. Electrophoresis separates charged molecules according to their size, shape and charge. Typically a gel made of either agarose or polyacrylamide provides the support system for resolving molecules in an electric field. The macromolecules are placed in a well at one end of the gel and an electrical current is applied through a buffer across the system. Gels made from polymerized acrylamide (polyacrylamide gels) are frequently used for separating proteins and small-sized nucleic acids because they have high resolving power, do not interact with biomolecules, and have a stable matrix. Typically the polyacrylamide is crosslinked with N,N’-methylene-bis acrylamide in a n aqueous 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 formed 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 or plastic that are sealed around the edges and are stood upright during polymerization. A comb is inserted at the top edge of a gel after pouring the polymerizing reaction between the plates, in order to produce the wells into which the samples are placed. 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 the same concentration throughout the gel or 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 polyacrylamide/bis-acrylamide and is poured first. On top, there is a stacking gel that contains a lover percentage of polyacrylamide/bis-acrylamide and prepared with a buffer containing a lower concentration of buffer salts. The stacking gel has lower conductance than the resolving gel. This means that the proteins will carry the current more and therefore travel quickly through the stacking gel until it reaches the resolving gel, where they concentrate at the interface, in a “stack”. This allows the proteins to enter the resolving gel close together and therefore increases the resolution during electrophoresis in the resolving gel, where the smaller pore size exerts a stronger frictional drag on the proteins during their separation.

Native gels

Native gels are run on proteins when the native 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 often performed with native gels to ensure that the antibody detection of a protein of interest will work.

When an electrical field is applied to the proteins they will travel from the wells where they were loaded, down into the acrylamide gel towards the positive electrode (the anode) IF the net charge of the protein is negative. While the electric field accelerates negatively charged proteins towards the anode, their rates of movement is influenced by frictional drag, in turn affected by factors such as size and shape of the molecule. 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 fibrous proteins having a more extended structure in an electrophoretic gel.

The ratio of charged amino acids in a protein influences movement through the gel by determining the net charge on the protein; 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 the wrong direction from the sample well, away from the anode and towards the cathode. The net charge of a protein can be manipulated by a change in pH. Buffers at lower ranges of pH will shift the dissociation of acidic amino acid side groups (aspartate and glutamate) towards their protonated, or uncharged state. At the same time these acidic buffers will shift the basic amino acid side groups (arginine, lysine, and histidine) towards their protonated, or positively charged state. If basic buffering systems are used, the acidic side groups will become deprotonated and acquire a negative charge, while the basic side groups will shift towards their deprotonated, or uncharged state. For that reason, most native gel buffering systems operate in a pH range of 8-9, where all but the most basic of proteins will carry a net negative charge and will migrate into the gel towards the anode.

The rate of migration in a native gel is not strictly a function of size, but is influenced by the size-to-mass ration of the proteins. Highly acidic proteins will have a higher negative charge per unit mass than a more basic protein, so will migrate faster during electrophoresis regardless of size differences between the two proteins.

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 relative size of proteins separated on a native gel. In order for proteins to behave similarly in a electric field, their differences in charge-to-mass ratio and their difference in shape must be made to be uniform. This uniformity is accomplished by denaturation of proteins into a uniformly extended shape and coating of proteins with negative charges. That way the unfolded proteins will be accelerated uniformly by the electric field and the frictional drag through the gel matrix will be a simple function of size.

The denaturation and coating of protein molecules with negative charges is accomplished by the use of heat and an ionic detergent called sodium dodecyl sulfate (SDS). This detergent is the sodium salt of a 12-carbon alkyl sulfate compound. In combination with heat, 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. Once denatured, SDS will coat the denatured proteins’ hydrophobic amino acid side groups with its hydrophobic dodecyl tail, thereby coating the protein with negative charges from the sulfate head group of the detergent. As long as SDS remains in solution with a denatured protein, it will not renature.

Reducing agents are often added to SDS in order to break any disulfide bonding that holds tertiary and quaternary proteins structures in a protein molecule. Dithiothreitol and 2-mercaptoethanol are short molecules containing a sulfhydryl (-SH) group that will convert the disulfide cystine bridges into two cysteine side groups to release 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.)

Typically, 1-2% SDS and 0.1 M mercaptoethanol buffered at pH 6.8 are used along with high temperatures to completely denature proteins and coat their extended structures with negative charges. In addition to 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 g protein are loaded into a well, depending on the purity of proteins in the sample. This total volume is typically about 20-40 L, half of which is protein and half is the 2X sample buffer.

Gels are generally run in a Tris-glycine buffer system with 0.1% SDS, pH 8.9. When the tracking dye from the sample buffer reaches the end of the resolving gel, proteins can be fixed into place (preventing diffusion of the separated bands) by acidified methanol solutions. The protein bands can be analyzed directly following a staining procedure with Coomassie blue dye. This gives rise to the SDS-PAGE electrophoresis that is the most common method of identifying an unknown protein or determining its molecular weight. A set of proteins markers of known molecular weight are run on the same gel 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.

Blotting Techniques

All blotting techniques use the same principle: macromolecules are transferred out of a gel onto another matrix, either nylon or nitrocellulose. This matrix may then be queried by a probe to visualize the bands of molecules. For example, in Southern blotting, DNA is subjected to electrophoresis and then a nylon sheet is used as the matrix to “blot” the gel. The nylon is treated with radioactively-labeled DNA probe molecules that will bind to specific bands on the membrane. The resulting DNA is then visualized by autoradiography. In a Western blot procedure, a specific protein separated on an electrophoretic gel can be identified by an immunological staining procedure. The separated proteins must be prepared for antibody binding by transfer to a membrane by electroelution, the “blotting” technique.

The following table summarizes the kinds of blotting that are typically done and the probes used.

Blotting system

/ Molecules to be identified / Probe
Southern / DNA / Radiolabelled
DNA
Northern / RNA
(usually mRNA) / Radiolabelled
DNA
Western / Protein / Antibody
South-Western / protein
(which can bind to DNA) / dsDNA

The steps of the Western blotting procedure for analysis of proteins is listed below:

1) Electrophoresis. Proteins are separated by gel electrophoresis, usually by SDS-PAGE using a tris/glycine/SDS running buffer.

2) Transfer. The proteins are transferred to a sheet of special membrane, usually nitrocellulose polyvinyl pyrolidon, or nylon, though other types of paper, or membranes, can be used.. This transfer is generally done by electroelution at 90 degrees to the gel through a buffer. The proteins retain the same pattern of separation they had on the gel. The blotting buffer is generally similar to the electrophoresis buffer, but lacking SDS. An alcohol (methanol, ethanol, or isopropanol) is added to the blotting buffer to facilitate protein binding to the membrane.

3) Protein Visualization.

a)Blocking. The blot is incubated with a generic protein (such as nonfat milk proteins) and a nonionic detergent such as Tween 20 to bind to any remaining sticky places on the nitrocellulose. This prevents nonspecific binding of antibody to the membrane.

b)Primary Antibody. An antibody (monoclonal or polyclonal) directed against the protein of interest is then incubated with the gel to allow it to bind to its specific protein. The antibody is diluted in a buffer solution, generally phosphate-buffered saline (PBS) containing a carrier protein, generally bovine serum albumin (BSA), along with some nonionic detergent such as Tween 20. The additives to the buffer help to ensure that the antibody is specific for the protein of interest, and does not bind to other proteins on the membrane.

c)Secondary Antibody. Since the antigen-antibody complexes are not colored, they must be treated in some way in order to visualize them. Usually an enzyme such as a horseradish peroxidase or alkaline phosphatase is coupled to a secondary antibody that binds to immunoglobulin (Ig) chains of the primary antibody. Alternatives to the conjugated enzyme are radiolabelling or dye conjugation of the secondary antibody.

d)Developing. The unbound secondary antibodies are washed away, and the conjugated enzyme is then presented with a colorless substrate which when reacted, will produce a colored product. The appearance of a colored enzyme product indicates the location on the gel of the protein of interest. Band densities in different lanes can be compared providing information on relative abundance of the target protein. Thus, the Western blot is similar to an ELISA. The major difference is the resolution and transfer to a membrane of the protein before being presented with antibodies.

The table below lists the enzymes and substrates commonly used for detection of target proteins in a Western blot.

Enzyme / Substrate / Color of reaction product
Alkaline phosphatase / 5-bromo-4-chloro-3’-indolylphosphate with nitroblue tetrazolium chloride (BCIP/NBT)
naphthol AS-MX phosphate with Fast Red TR / Black/purple
Bright red
Horseradish peroxidase / 4-chloronaphthol (4-CN)
3-amino-9-ethyl carbazole (AEC)
3,3’-diaminobenzidene tetrahydrochloride (DAB)
(note: carcinogenic!) / Blue/purple
Red/brown
Brown
Glucose oxidase / Phenazine methosulfate with nitroblue tetrazolium chloride / Black/purple

Troubleshooting and optimizing a Western blot

Like all complex procedures, there are many technical difficulties that can lead to disastrous results. Careful attention to the correct temperatures and incubation times is very important to the success of a Western blot, as is the quality of the reagents being used. The blotting step itself is critical: it is important to layer the gel onto the onto a prewetted membrane with good contact and without any air bubbles. The elution time must be optimized for the size of protein of interest. Overheating during electroelution must be carefully avoided for efficient transfer of proteins to a membrane. The following guide can help to be used to optimize a Western blot procedure for a specific application, and to determine exactly which step of the protocol may be a problem when getting disappointing results.

Problems associated with the electrotransfer blotting stage:

Symptom / Possible cause / Remedy
Band smeared/distorted / Membrane not uniformly wetted prior to transfer.
Air bubbles under membrane and between other layers in the stack.
Uneven contact between gel and membrane
Too much heat generated during the transfer
Proteins transferred too rapidly; protein buildup on the membrane surface / Many types of membranes are hydrophobic, and must be prewetted with methanol; the entire membrane should change uniformly from opaque to semi-transparent.
Using a pipet or stirring rod, gently roll out any trapped air bubbles while assembling the stack.
Make sure the entire gel and membrane surfaces are in good contact.
Pre-chill the buffer, carry out the transfer in a cold room, or reduce the current.
Reduce the strength of the electrical field.
Weak signal / Incomplete transfer of proteins
Proteins passing through the membrane.
Proteins retained in the gel.
Isoelectric point of the protein is at or close to the pH of the transfer buffer.
Poor protein retention on membrane / Stain the gel after the transfer to check for residual proteins. If transfer was not complete, review your transfer technique. Improper buffer concentrations, or too much methanol, for example, can lead to poor transfer efficiency.
Increase the time the proteins have to interact with membrane during the transfer by reducing the voltage by as much as 50%.
Highly negatively charged proteins tend to move quickly in an electric field. Decrease the voltage to slow down migration of these proteins
The presence of SDS in the gel may inhibit protein binding. Equilibrate the gel in the transfer buffer for at least 15 minutes prior to the transfer.
Methanol concentration in transfer buffer is too low to facilitate removal of SDS. Increase the methanol to 15-20%, especially for smaller molecular weight proteins.
Inadequate prewetting of the membrane will impede binding of proteins.
If the methanol concentration in the transfer buffer is too high, it can remove SDS from proteins and lead to protein precipitation in the gel, reducing the transfer of large proteins out of the gel. If this is an issue, the transfer buffer can be supplemented with SDS (0.01%-0.05%) or methanol concentration can be reduced.
A protein has no net charge at a pH that matches its isoelectric point, so will not migrate in an electric field. Try a higher pH buffer such as 10mM CAPS buffer at pH 11, including 10% methanol.
Once the transfer is complete, be sure to dry the membrane completely to obtain optimal binding and fixation of the proteins. This should be done prior to any downstream detection method.
No signal / No transfer of proteins / Check for the gel and membrane orientation during the transfer process. Use pre-stained molecular weight standards to monitor the transfer.
Poor transfer of small sized proteins (<10kDa) / SDS interferes with binding of small molecular weight proteins.
Low methanol concentration in the transfer buffer.
Insufficient protein binding time.
Current doesn’t pass through the membrane. / Remove SDS from the transfer solution.
Use higher % methanol (15-20%) in the transfer buffer.
A lower voltage may optimize binding of small proteins to the membrane.
Cut the membrane and blotting paper exactly to the gel size; do not allow overhangs.
Poor transfer of large sized proteins (>80 kDa) / Methanol concentration is too high. / Reducing the methanol concentration to 10% (v/v) or less should help in the transfer process by allowing the gel to swell and reducing SDS loss, thereby reducing the protein precipitation in the gel. Larger proteins may require a longer transfer duration or a higher current setting.
Poor transfer of a wide range of protein sizes. / Different conditions required to transfer large and small proteins / Try this protocol: “Transfer of a brad MW range of proteins may require a multi-step transfer” by t. Otter et al, Anal. Biochem. 162:370-377 (1987)

Problems associated with the protein visualization stage: