Analysis of Gene Expression at the Proteomic Level


Introduction Principles

Extraction of Cellular Proteins

Method A. Extraction of Total Proteins Using Lauroylsarcosine Buffer

Protocol 1. Protein Extraction from Cultured Cells Protocol 2. Protein Extraction from Tissues Analysis of Proteins by SDS-PAGE and Western Blotting Protocol 1. Preparation of the SDS Separation Gels Protocol 2. Preparation of Stacking Gels Protocol 3. Electrophoresis

Protocol 4. Staining and Destaining of SDS-PAGE Using a Modified Coomassie Blue (CB) Method (Figure 11.2)

Protocol 5. Transfer of Proteins from SDS-PAGE onto a Nitrocellulose or PVDF Membrane by Electroblotting (Figure 11.3) Protocol 6. Blocking and Hybridization of the Membrane Filter Using Specific Antibodies

Protocol 7. Detection of Hybridized Signals Analysis of Proteins by 2-D Gel Electrophoresis Troubleshooting Guide


Proteins are translated from mRNA species and considered to be the final products of gene expression. It is the protein that facilitates the specific function of a gene that makes gene expression meaningful.1,2 Therefore, protein expression is one of the most important parts of modern molecular biology. To explore proteins, one key approach is to analyze them by electrophoresis, a process in which a net charged molecule will move in an electric field. SDS-PAGE (sodium dodecyl sulfate poly-acrylamide gel electrophoresis), which was developed in the mid-1960s, is a widely


used technique to separate proteins according to their net charges, sizes and shapes.13-6 To identify a specific protein from a protein mixture, it may be necessary to use a powerful and commonly employed technique: western blotting.

Western blot hybridization or immunoblotting is a procedure in which different types of proteins are separated by SDS-PAGE and immobilized onto a solid support PVDF (polyvinylidene difluoride) or nitrocellulose membrane. The protein of interest on the blotted membrane is then detected by incubating the membrane with a specific antibody as a probe. The name "western" is not a person's name, nor is it due to a geographic location. This technique was developed later than the Southern blot method and thus named western blotting. Because the technique mandates specific antibodies, (monoclonal or polyclonal), it is also called immunoblotting. Western blot technique is a sensitive, reliable and quantitative method widely employed in analysis of proteins. It can simultaneously offer information about the species, sizes and expression levels of diverse proteins that cannot be obtained by other alternative techniques.1,2,4-8

General factors should be considered when western blot hybridization is carried out. The concentration of polyacrylamide gel will influence separation of proteins. Depending on the particular size of the protein of interest, a general chart is displayed below. Normally, the larger the size is of a protein to be detected, the lower the percentage of polyacrylamide gel that should be used in order to obtain a sharp band.

During SDS-PAGE, proteins move through the gel matrix at a rate inversely proportional to the log of their molecular weight. Small proteins migrate faster than large molecules.

Transfer of proteins from a polyacrylamide gel onto a solid nitrocellulose or nylon membrane can be completed by the electrophoretic blot method. Compared with traditional nitrocellulose membranes, PVDF membranes are durable and tear resistant. Based on our experience, we highly recommend that PVDF be used as the solid transfer support.

Western blotting requires specific antibodies as probes. The specificity and activity of an antibody are absolutely essential to the success of western blot hybridization. Two general types of antibodies are monoclonal antibodies raised in mice, and polyclonal antibodies usually raised in rabbits.4 A monoclonal antibody has a single antibody specificity, a single affinity and a single immunoglobulin isotype. However, a preparation of polyclonal antibodies contains a variety of antibodies directed against the antigen of interest as well as nonspecific proteins. As a result, a monoclonal antibody is more specific than polyclonal antibodies. Monoclonal antibodies are produced by a monoclonal population of cells derived from one cloned cell. Hence, all the antibodies are theoretically identical.

Polyacrylamid % (w/v)

Protein size (kDa)

FIGURE 11.1 Scheme of detection of specific protein by western blot hybridization.

A standard western blot hybridizaiton mainly consists of four procedures: separation of proteins by SDS-PAGE, transfer of proteins onto a solid membrane, incubation of the membrane with specific antibodies and detection of hybridized signals (Figure 11.1). This chapter describes step-by-step protocols for successful western blotting.


Knowing the principles of western blotting is extremely important for success. The primary principles of western blotting or immunoblotting start with separation of proteins by SDS-PAGE. SDS is a strong, negatively charged detergent composed of a hydrophilic head and a long hydrophobic tail (CH3-CH2-CH2-CH2-CH2-CH2-CH2-CH2-CH2-CH2-CH2-CH2-SO4Na+). SDS serves to denature proteins and make them negatively charged. To better understand how a protein molecule is denatured by detergents, it is necessary to review the structures of proteins briefly.

In general, functional proteins have four levels of structures. Primary structure is the amino acid sequence that is the backbone of a protein. Linear polypeptide chains undergo appropriate folding into the secondary structure of proteins by hydrogen bonds between main-chain NH and CO groups. In some proteins, disulfide bridges or bonds (-S-S-) are formed between the sulfhydryl groups of two cysteine residues. The tertiary structure of protein architecture refers to the spatial folding following the secondary structure. Tertiary structure is the highest level of structure for most proteins. However, many proteins contain multiple peptide chains or sub-units. The spatial arrangement of such subunits and the nature of their connections are called quaternary structure. Proteins with multiple subunits mandate the quaternary structure for functions. These unique structures make proteins much more stable. However, for SDS-PAGE analysis of proteins, proteins need to be disrupted to their primary structures using appropriate detergents. SDS is an anionic detergent that binds to hydrophobic regions of protein molecules and causes them to unfold into extended polypeptide chains. As a result, the individual proteins are dissociated from other proteins and rendered freely soluble in the SDS solution. In addition, a reducing agent, b-mercaptoethanol (SH-CH2-CH2-OH), is commonly used to break any S-S bonds in proteins.

During SDS-PAGE, the anions of SDS not only denature proteins but also coat polypeptides at a ratio of about one SDS for every two amino acid residues, which gives approximately equal charge densities per unit length and makes the polypep-tides negatively charged. As a result, each protein molecule binds large numbers of the negatively charged SDS molecules that overwhelm the protein's intrinsic charge. This is one of the principles of protein separation by SDS-PAGE. In an electric field, proteins wrapped with the negative SDS migrate toward the positive electrode. To enhance separation, polyacrylamide gels or PAGEs are widely used as supporting media and molecular sieves. The meshes and pore size of a PAGE can be readily made by the polymerization of acrylamide and a cross-linking reagent methyleneb-isacrylamide using appropriate concentrations. When a voltage is applied, proteins of the same size have the same amount of SDS molecules and the same shape due to complete unfolding by the SDS, and thus migrate at the same speed in a given pore size of a slab SDS-PAGE.

Smaller proteins move faster than larger proteins, which are retarded much more severely. As a result, proteins are fractionated into a series of discrete protein bands in order of their molecular weight (MW). There is a linear relationship between the log of the MW of a polypeptide and its Rf. Rf is the ratio of the distance from the top of the gel to a band divided by the distance from the top of the gel to the dye front. A standard curve is generated by plotting the Rf of each standard polypeptide or band marker as the abscissa and the log10 of its MW as the ordinate. The MW of an unknown protein band can then be determined by finding its Rf that vertically crosses on the standard curve and reading the log10 MW horizontally crossing to the ordinate. The antilog of the log10 MW will be the actual MW of the protein. Following electrophoresis, the SDS-PAGE gel can be stained with Coomassie blue solution. The major proteins or bands are then readily displayed by destaining the gel (Figure 11.2).

Inasmuch as the separated proteins are coated with negatively charged SDS, they can be readily transferred from the SDS-PAGE onto a solid membrane by electroblotting. The negatively charged proteins migrate toward the positive electrode and are retained on the membrane placed between the gel and the positive electrode (Figure 11.3). Various bands at different positions are exactly blotted at appropriate positions on the membrane. To check the efficiency of blotting, the blotted membrane can be stained by Ponceau S solution and major bands are readily seen if the transfer is successful.

The major principle of western blotting is the interaction between a specific antibody and its target protein on the membrane (Figure 11.4). To understand how the binding of antibody-antigen occurs, it is helpful to be briefly familiar with

FIGURE 11.2 Coomassie blue staining and destaining of SDS-PAGE. Lane 1 to lane 3 represent proteins extracted from three different cell lines.

FIGURE 11.2 Coomassie blue staining and destaining of SDS-PAGE. Lane 1 to lane 3 represent proteins extracted from three different cell lines.

Anode paper sheets

FIGURE 11.3 Diagram of semidry electric blotting of proteins onto a membrane.

Anode paper sheets

FIGURE 11.3 Diagram of semidry electric blotting of proteins onto a membrane.

structures of antibodies. An antibody is a Y-shaped molecule containing two heavy chains, two light chains and two identical antigen-binding sites or bivalent at the tip of two arms. It is important to know that the binding sites are specific for a particular antigen (e.g., a protein). The antigen-binding sites make it possible for an antibody to cross-link the antigenic determinants of the target antigen or a protein in terms of western blotting or immunoblotting. Many antigens have multiple antigenic determinants or epitopes, which enable the antigen to form linear chains or cyclic complexes with the antibody.

The antibody that directly binds to its antigen is called the primary or first antibody. In western blot hybridization, a secondary antibody is needed, which usually interacts with the primary one (Figure 11.4). The secondary antibody is

frequently conjugated with an enzyme such as alkaline phosphatase or horseradish peroxidase. Once a complex of protein-primary antibody-secondary antibody is formed on a blotted membrane, it can be detected using substrates such as NBT and BCIP for alkaline phosphatase or luminol for horseradish peroxidase (Figure 11.5).

FIGURE 11.5 Detection of actin from the proteins extracted from mouse embryonic stem cells. Monoclonal antibodies against actin were used in western blotting.


Method A. Extraction of Total Proteins Using Lauroylsarcosine Buffer

Traditionally, proteins are extracted by lysing cells with SDS gel sample loading buffer followed by boiling. This procedure rapidly inactivates proteolytic enzymes and the protein samples can be immediately used for sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) or two dimensional (2D) gel electrophoresis; however, they are not suitable for quantitative analysis of the samples' protein concentration because the samples contain SDS and 2-mercaptomethanol. In the Bradford assay, these compounds significantly decrease the O.D. at A595 nm, so the concentration of proteins in the extracts cannot be accurately measured. An alternative procedure is to determine the concentration of the proteins by the method of the bicinchoninic acid (BCA) assay. The major drawbacks of this approach are that these procedures are time consuming, and a portion of proteins are lost during the process of trichloroacetic acid (TCA) precipitation as well as centrifugation. As a result, the protein concentration determined by this method is lower than what truly exists in the sample.

To solve the problem, a new extraction buffer has been developed.4 The proteins extracted are suitable for quantitative analysis using the Bradford assay and for routine analysis of proteins by SDS-PAGE, 2-D gel electrophoresis, and immunob-lotting. This method provides an economical, rapid, simple and efficient approach for protein analysis. Cells in tissue culture dishes or flasks can be directly lysed with an extraction buffer containing 45 mM Tris-HCl (pH 6.8), 0.2% ^-Lauroylsarcosine (sodium salt, Sigma Chemical Co., and 0.2 mM of phenylmethanesulfonyl fluoride (PMSF). Because of the instability of PMSF in aqueous solution, it is recommended to add this component to the buffer shortly before use.

Protocol 1. Protein Extraction from Cultured Cells

1. When the cells reach 50 to 100% confluence, aspirate the medium and add an appropriate volume of extraction buffer (1 to 1.5 ml/100-mm dish).

2. To lyse the cells, incubate for 5 min at room temperature with occasional agitation by hand. To confirm cell lysis, the dish or flask containing the cells can be observed by phase contrast microscopy.

3. Transfer the lysed mixture into a microcentrifuge tube (Eppendorf) and store at -20°C until use. At this point, the sample can be utilized, without further processing, for protein concentration measurement by the Bradford assay and for quantitative analysis of proteins by 1D or 2D SDS-PAGE and immunoblotting.

Protocol 2. Protein Extraction from Tissues

1. Completely homogenize 1 g tissues of interest in 2 ml of extraction buffer.

2. To lyse the cells, incubate the homogenate for 10 min at room temperature with occasional agitation by hand.

3. Centrifuge at 10,000 x g for 2 min and carefully transfer the supernatant into a microcentrifuge tube and store at -20°C until use. At this point, the sample can be utilized, without further processing, for protein concentration measurement by the Bradford assay and for quantitative analysis of proteins by 1D or 2D SDS-PAGE and immunoblotting.

Method B. Determination of Protein Concentration Using the Bradford Assay

1. Label an appropriate number of cuvettes for standard curve and protein samples.

2. To the cuvettes used for standard curve, add the following components in the order shown.

a. BSA stock solution (1 mg/ml in dH2O): 0, 5, 10, 15, 20, 25, 30 to 40 ml, respectively.

b. The protein extraction buffer: 20 to 40 ml for every cuvette, depending on the volume of protein sample used for measurement.

c. dH2O: add an appropriate volume to each cuvette to give a total volume of 0.1 ml.

d. Bradford reagent solution: 0.9 ml to every cuvette, producing a total volume of 1 ml.

3. Transfer each protein sample (20 to 40 ml) extracted from cultured cells or from tissues to a plastic cuvette. Add dd.H2O to 0.1 ml, and then 0.9 ml of the Bradford reagent solution.

4. Carry out protein measurement at a wavelength of 595 nm in a UV-Vis spectrophotometer according to the manufacturer's instructions.


Protocol 1. Preparation of the SDS Separation Gels

1. Thoroughly clean the glass plates and spacers with detergent and wash them in tap water. Rinse the plates and spacers with dd.H2O several times and air-dry.

2. Wear gloves and, using clean paper prewet with 100% ethanol, wipe dry the glass plates and spacers. Assemble the vertical slab gel unit such as the Protein II (BioRad) or the SE 600 vertical slab gel unit (Hoeffer) in the casting mode according to the instructions. Briefly, place one spacer (1.5 mm thick) on each side between the two glass plates and fix in place with clamps, forming a sandwich. Prior to casting the sandwich tightly, vertically place the sandwich on a very level bench for an appropriate adjustment. Make sure that the glass plates and spacers are well matched to prevent any potential leaking. Repeat for the second sandwich if needed. Tightly cast the sandwich in the casting mode.

Note: The glass plates should be in very good shape with no damage at the edges. If necessary, spray a little grease or oil on the spacer areas at the top and bottom ends of the sandwich to prevent leaking.

3. Test potential leaking by filling an appropriate volume of dd.H2O into the sandwich. Drain away the water by inverting the unit.

4. Prepare the separation gel mixture in a clean 100-ml beaker or flask in the order shown. This is sufficient for two pieces of standard size gel: 20 ml of 30% monomer solution

15 ml of separating gel buffer 0.6 ml of 10% SDS solution

24.1 ml of dd.H2O Mix after each addition.

Caution: Acrylamide is neurotoxic. Gloves should be worn when handling the gel mixture.

5. Add 0.25 ml of freshly prepared 10% ammonium presulfate (AP) solution and 20 ml TEMED to the gel mixture. Briefly mix by gently swirling or using a pipette.

6. Immediately and carefully fill the mixture into the assembled sandwich up to about 4 cm from the top edge using a 10- to 15 ml-pipette or a 50ml syringe.

7. Take up approximately 1 ml of dd.H2O into a 1-ml syringe equipped with a 22-gauge needle and carefully load 0.5 ml of the water, starting from one top corner near the spacer, onto the surface of the acrylamide gel mixture. Load 0.5 ml of the water from the other top corner onto the gel mixture. The water will overlay the surface of the gel mixture to make the surface very even. Allow the gel to be polymerized for about 30 min at room temperature. A very sharp gel-water interface can be seen once the gel has polymerized.

8. Drain away the water layer by inverting the casting unit and rinse the surface once with 1 ml of gel overlay solution. Replace the solution with 1.5 ml of fresh gel overlay solution for at least 5 min. The gel can be kept at this point for 2 h to overnight prior to the next step.

Protocol 2. Preparation of Stacking Gels

1. Prepare the stacking gel mixture in a clean 50-ml beaker or a 50-ml flask as follows:

2.66 ml of 30% monomer solution 5 ml of stacking gel buffer 0.2 ml of 10% SDS

12.2 ml of dd.H2O Mix after each addition.

2. Drain away the overlay solution on the separating gel by tilting the casted unit and rinse the surface with 1 ml of the stacking gel mixture prepared at the previous step. Drain away the stacking gel mixture and insert a clean comb (1.5 mm thick with 14 or 30 wells) from the top into the glass sandwich.

3. Add 98 ml of 10% AP solution and 10 ml of TEMED to the stacking gel mixture. Gently mix and add the gel mixture into the sandwich from both upper corners next to the spacers using a pipette or an equivalent.

Note: The stacking gel mixture may be filled into the sandwich prior to inserting the comb. However, air bubbles are readily generated and trapped around the teeth of the comb. It is recommended that the comb be inserted into the sandwich prior to filling the stacking gel mixture.

4. Allow the stacking gel to polymerize. It usually takes approximately 30 min at room temperature.

Protocol 3. Electrophoresis

Loading Protein Samples and Protein Standard Markers onto the Gel

1. While the stacking gel is polymerizing, prepare samples and standard markers. To denature proteins completely, add one volume of 2X sample loading buffer to each sample and standard marker in individual microcentrifuge tubes. Tightly cap the tubes and place them in boiling water for 4 min. Briefly spin down.

Note: The loading buffer contains SDS and mercaptoethanol, which are strong detergents for denaturation of proteins. The amount of proteins loaded into one well should be 5 to 30 fig for Coomassie blue staining and western blotting. The amount of protein standard markers for one well should be 2 to 10 fig.

2. Carefully pull the comb straight up from the gel, and rinse every well with running buffer using a pipette. Drain away the buffer by tilting the cast sandwich. Repeat rinsing once.

3. If the apparatus Protein II (BioRad) is used, the sandwich can be directly cast in place according to the manufacturer's instructions. An upper buffer chamber will be readily formed. If the SE 600 Vertical Slab Gel Unit (Hoeffer) is utilized, carefully cast the upper buffer chamber in place according to the instructions. Remove lower cams and cam the sandwich tightly to the bottom of the upper buffer chamber.

Note: Apply a little grease or oil on the spacer areas at the top corners of the sandwich to prevent leaking.

4. Fill the upper chamber with an appropriate volume of running buffer.

5. Take up sample or markers into a pipette tip or a syringe equipped with a 22-gauge needle. Insert the tip or needle into the top of a well and load the sample into it. Because of the 2X loading buffer added, the sample is heavier than running buffer and can be seen sinking to the bottom of well when loading.

Note: The volume for each well should be less than 40 ml for a standard gel or less than 10 ml for a minigel. Overloading may cause samples to float out, resulting in contamination among wells. The markers should be loaded into the very left or right well or both wells.

Carrying Out Electrophoresis

1. Carefully insert the assembled unit with loaded samples into the electro-phoresis tank filled with 2 to 4 l of running buffer. Remove any bubbles trapped at the bottom of the sandwich using a glass Pasteur pipette hook or an equivalent tool.

Note: The volume of running buffer in the tank should be sufficient to cover two thirds of the sandwich. Otherwise, the heat generated during electro-phoresis will not be uniformly distributed, causing distortion of the band patterns in the gel.

2. Place the lid, or cover, on the apparatus and connect to a power supply. The cathode (negative electrode) should be connected to the upper buffer chamber and the anode (positive electrode) must be connected to the bottom buffer reservoir. Proteins negatively charged by SDS will migrate from the cathode to the anode.

3. Apply a constant power or current at 25 to 30 mA/1.5 mm thick standard size gel. Allow the gel to run for several hours or until the dye reaches about 1 cm from the bottom of the gel. If an overnight run is desired, the current should be adjusted empirically.

4. Turn off the power and disconnect the power cables. Loosen and remove the clamps from the sandwiches. Place the gel sandwiches on the table or bench. Carefully remove the spacers and separate the two glass plates starting from one corner by inserting a spacer into the sandwich and simultaneously lifting the top glass plate. Using a razor blade, make a small trim at the upper left or right corner of the gel to record the orientation. One gel will be stained to determine the MW and patterns of different protein bands. The other gel will be used for western blotting.

Protocol 4. Staining and Destaining of SDS-PAGE Using a

Modified Coomassie Blue (CB) Method (Figure 11.2)

1. Wear gloves and carefully transfer one of the gels into a glass or plastic tray containing 100 to 150 ml of CB staining solution and allow the gel to be stained for 30 min in a 50°C water bath with occasional agitation.

2. Replace the staining solution with 150 ml of rapid destaining solution. Allow destaining to take place for 60 to 80 min on a shaker with slow shaking. Change the solution once every 20 min.

Notes: (1) Major protein bands should be visible after 20 min of destaining; all patterns of bands with a clear background will appear following 80 min of destaining. If desired, the destained gel can be photographed or dried or kept in 50% diluted destaining solution in distilled water for up to 1 month. (2) The gel may be rapidly destained using 1 to 1.5% bleach solution in distilled water. Allow destaining to proceed for 30 min with agitating. Constantly monitor the destaining and immediately rinse the gel with a large volume of top water once clear bands are visible. Change distilled water several times until the blue color of the bands is stable. The drawback to this procedure is that, if overly destained, some bands may disappear. (3) The gel can be stained by the silver method, which is more sensitive than Coomassie blue staining. Some weak bands seen in Coomassie blue staining can become sharp bands after silver staining. However, staining and destaining procedures are quite complicated and time consuming, so silver staining is not considered to be within the scope of this lab protocol.

3. To determine the MW of specific bands accurately, measure the distance from the top of the gel to each of the bands in the protein standard marker lane and to specific bands in the protein sample lanes. Calculate the Rf for each band: divide the distance from the top of the gel to a specific band by the distance from the top of the gel to the dye front. Make a standard curve by plotting the Rf of each standard marker band as the abscissa and the log10 of each MW as the ordinate. The MW of an unknown band in the protein sample can readily be determined by finding its Rf that vertically crosses the standard curve and the log10 MW that horizontally crosses to the ordinate. The antilog of the log10 MW is the actual MW of the protein band.

Note: If accurate determination of protein bands is not necessary, the MW of an unknown band can be estimated by comparing the position of a specific band with those of standard marker bands.

Protocol 5. Transfer of Proteins from SDS-PAGE onto a Nitrocellulose or PVDF Membrane by Electroblotting (Figure 11.3)

Method A. Semidry Electroblotting

1. Following SDS-PAGE, soak the gel in 200 ml of transfer buffer for 10 min.

2. Cut a piece of polyvinylidene difluoride (PVDF) or nitrocellulose membrane and 10 sheets of Whatman filter paper to the same size as the gel and soak them in transfer buffer for 4 min prior to blotting. If PVDF is used, briefly soak the membrane in methanol for 30 s prior to soaking in transfer buffer.

3. Assemble a blotting system in the order shown in Figure 11.3 using a Multiphor II electrotransfer apparatus (Pharmacia Biotech).

a. Rinse the bottom plate (the negative electrode) in distilled water and place five sheets, one by one, of the soaked 3MM Whatman paper on the plate. Remove any bubbles underneath by rolling a pipette over the filter papers.

b. Place the soaked membrane on the top of 3MM Whatman sheets. Gently press the membrane to remove any bubbles underneath.

c. Carefully overlay the membrane with the gel, starting at one side of the gel, then slowly proceed to the other end. The marked side of the membrane should face the gel. Carefully remove any bubbles between the membrane and the gel, which can be done by gently pressing or rolling on the membrane.

d. Gently overlay the gel with five sheets, one by one, of the presoaked 3MM Whatman paper.

e. Prewet the top plate (the positive electrode) in distilled water and carefully place the plate on top of the assembled unit.

To avoid producing bubbles, do not disturb the assembled unit lying underneath.

f. Place a bottle or beaker containing approximately 1000 ml water on the top plate to serve as a weight.

4. Connect the assembled unit to a power supply (positive to positive pole, negative to negative pole). Turn on the power and set it at a constant current. Allow blotting to proceed for 1 h at 1.2 mA/cm2 membrane at room temperature.

Notes: (1) If several gels need to be transferred simultaneously, place four sheets of 3MM Whatman paper between each gel and membrane, and adjust the current accordingly. (2) Air bubbles should be removed from each layer of the blot by carefully rolling a pipette over the surface. It is important not to move gel, membrane and blotting paper until transfer is complete. Otherwise, bubbles will be generated and band positions are likely to be changed.

Method B. Liquid Transfer Using a Hoeffer TE 42 Blotting Apparatus

1. Soak the gel, membrane and four pieces of 3MM Whatman paper sheets (relatively bigger than the gel) in 500 ml of transfer buffer for 15 min.

2. Fill a tray large enough to hold the cassette with transfer buffer to a depth of approximate 4 cm.

3. Place one half of the cassette in the tray with the hook facing up and put one DacronTM sponge on the cassette.

4. Place two sheets of the soaked 3MM Whatman paper on the sponge and remove any bubbles.

5. Overlay the soaked membrane on the paper sheets and remove any air bubbles between the layers.

6. Carefully place the soaked gel on the membrane and overlay the gel with two sheets of presoaked 3MM Whatman paper.

Note: Gloves should be worn. Gently press the filter papers to force out any trapped air bubbles that will locally block transfer of proteins onto the membrane.

7. Overlay another Dacron sponge on the filter sheets and place the second half of the cassette on the top of the stack.

8. Hook the two halves together by sliding them toward each other so that the hooks are engaged with the opposite half.

9. Place the assembled cassette into the blotting chamber according to the instructions. Fill the chamber with transfer buffer sufficient to cover the cassette. Place a stir bar in the chamber.

Note: It is extremely important to insert the cassette in the right place so that the membrane is between the gel and the anode (positive electrode).

10. Place the whole chamber on a magnetic stirrer plate and turn on the stir bar at low speed. Connect the apparatus to the power supply, turn on the power and set the current at 0.8 to 1 A.

11. Allow the transfer to take place for 60 to 75 min at 1 to 1.5 A.

Note: Because heat will be rapidly generated, the current should be monitored to be 1 to 1.5 amps. Otherwise, it may burn the apparatus.

Protocol 6. Blocking and Hybridization of the Membrane Filter Using Specific Antibodies

1. While the transfer is taking place, prepare blocking solution (see Materials Needed).

2. After transfer is complete, stain the membrane in 1X Ponceau S solution (Sigma Chemicals) diluted in distilled water using 10 ml per 12 x 14 cm2 membrane. Allow staining to take place for 5 to 8 min at room temperature and rinse the membrane in distilled water several times.

Note: A successful transfer should show red bands. Staining will not interfere with the following steps.

3. After rinsing, block the nonspecific binding sites on the membrane in blocking solution (100 ml per 12 x 14 cm2 membrane) at room temperature for 50 min with slow shaking.

Note: The membrane at this stage may be kept in the blocking solution for up to 48 h at 4 °C. To check the efficiency of transferring, the blotted gel may be stained with Coomassie blue and then destained. A successful blotting should have no visible bands left in the gel.

4. Replace the blocking solution with antibody solution containing primary antibodies (monoclonal or polyclonal antibodies against the protein of interest) and appropriate secondary antibodies diluted in PBS buffer. Allow incubation to proceed at room temperature for 70 to 100 min with slow shaking.

Please pay attention to the following notes: (1) Traditionally, the membrane is separately incubated with primary antibodies, washed several times and then incubated with secondary antibodies. In our experience, the membrane can be simultaneously incubated with first and second antibodies with high efficiency of hybridization. This saves at least 1.5 h. (2) The concentration of antibodies varies with their activity and specificity. Therefore, different dilutions, in case of new antibodies, must be tested in order to obtain the optimal concentration for hybridization. (3) If the primary antibodies are monoclonal antibodies from mice, the secondary antibodies should be commercial goat antimouse IgA, IgG, and IgM conjugated with alkaline phosphatase or with peroxidase or biotin labeled. If the primary antibodies are polyclonal antibodies from rabbits, the secondary antibodies should be goat antirabbit IgG conjugated with alkaline phosphatase or with peroxidase or biotin labeled. (4) The standard volume of antibody solution should be 30 to 40 ml per 12 x 14 cm2 membrane.

5. Transfer the hybridized membrane to a clean tray containing 200 ml/membrane of PBS. Allow washing to proceed for 60 min with slow shaking. Change PBS once every 20 min. Proceed to detection of hybridized signals.

Note: (1) The antibody solution can be stored at 4°C and be reused three to four times. (2) It is optional to add 0.05% (v/v) Tween-20 to PBS to wash the membrane.

Protocol 7. Detection of Hybridized Signals

Method A. Chemiluminescent Detection for Peroxidase-Conjugated or Biotin-Labeled Antibodies

1. Place a piece of SaranWrap film on a bench and add 0.8 ml/filter (12 x 14 cm2) of ECL detection solution (Amersham Life Science) or an equivalent to the center of the film.

2. Wear gloves and briefly dampen the filter to remove excess washing solution; thoroughly wet the protein-binding side by lifting and overlaying the filter with the solution several times.

3. Wrap the filter with SaranWrap film, leaving two ends of the film unfolded. Place the wrapped filter on a paper towel and, using another piece of paper towel, carefully press it to wipe out excess detection solution through the unfolded ends of the film. Excess detection solution will most likely cause a high background.

4. Completely wrap the filter and place it in an exposure cassette with protein-binding side facing up. Tape the four corners of the filter.

5. In a darkroom with a safe light on, overlay the filter with an x-ray film and close the cassette. Allow exposure to proceed at room temperature for 10 s to 15 h, depending on the intensity of the detected signal.

6. In a darkroom, develop and fix the film in an appropriate developer and fixer, respectively. If an automatic x-ray processor is available, development, fixation, washing and drying of the film can be completed in 2 min. If a hybridized signal is detected, it appears as a black band on the film (Figure 11.5).

Note: Multiple films may need to be exposed and processed until the signals are desirable. Exposure for more than 4 h may generate a high black background. In our experience, good hybridization and detection should display sharp bands within 1.5 h. In addition, the film should be slightly overexposed to obtain a relatively dark background that will help identify the sizes of the bands compared with the marker bands.

Method B. Colorimetric Detection for Alkaline Phosphatase-Conjugated Antibodies

1. Place the filter in color developing solution (10 ml/filter, 12 x 14 cm2) containing 40 ml of NBT stock solution and 30 ml of BCIP stock solution. NBT and BCIP stock solutions are commercially available.

2. Allow color to develop in a dark place at room temperature for 15 to 120 min or until a desired detection is obtained. Positive signals should appear as a blue/purple color.

Congratulations on your successful western blot hydribidation!

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