Western Blot Introduction

Western Blotting is an analytical Immunoblotting Technique to detect specific proteins in a cell extract or tissue homogenate. Western Blotting relies on the specific binding between the protein-of-interest and an antibody raised against this particular protein.

Step-by-step western blot protocol

All steps are carried out at room temperature unless otherwise indicated. Recipes for all standardized solutions highlighted in bold can be found on the internet or the back of a textbook.

SDS PAGE

• Construct an SDS-PAGE gel according to the molecular weight (MW) of your target protein(s). (Recommendations and gel recipes are presented at the end of this protocol)

• Tip 1: Tris-tricine gels separate low MW proteins (<20 kDa) better than Tris-glycine gels.

• Prepare samples in microfuge tubes. Add 4X SDS sample buffer so the total protein amount is 30–50 μg per sample (according to the protein amount measured by Bradford or BCA protein assay).
• Flick microfuge tubes to mix samples, and then heat to 95-100°C for 5 minutes.
• Set up electrophoresis apparatus and immerse in a 1x running buffer. Remove gel combs and cleanse wells of any residual stacking gel.
• Load samples and protein markers onto the gel using gel loading tips. Set electrophoresis power pack to 80V (through the stacking gel), before increasing it to 120V when the protein front reaches the separation gel.

Protein transfer

• PVDF membranes (or PSQ membranes with 0.22 μm micropores when MW of the target is <30 kDa) are strongly recommended. Soak membranes in methanol for 30 seconds before moving to transfer buffer. Soak the filter papers and sponges in the transfer buffer as well.
• Sequentially assemble the transfer constituents according to the illustration on page 7 of this booklet and ensure no bubbles lie between any of the layers. Apply semi-dry or wet transfer systems according to the manufacturer’s instructions.

Immunoblotting

• After transfer, wash the membrane twice with distilled water, and using a pencil, mark bands of the MW ladder on the membrane. If desired, stain the membrane with commercial Ponceau red solution for 1 min to visualize protein bands, then wash any Ponceau red staining with copious amounts of 1x TBST.
• Block with 1x TBST containing (2-5%) nonfat dry milk (or 1-5% BSA for the detection of phospho-epitope antibodies) with constant rocking for 1 hour or overnight at 4°C.
• Dilute primary antibody in blocking solution with a starting dilution ratio of 1:1000. (Optimal dilutions should be determined experimentally.) Incubate the membrane with primary antibody for 1 hour at room temperature, or overnight at 4°C.
• Wash membrane three times with 1x TBST for 10 minutes each.
• Incubate the membrane with a suitable HRP-conjugated secondary antibody (recognizing the host species of the primary antibody), diluted at 1:5000–1:50000 in blocking solution. Incubate for 1 hour with constant rocking.
• Wash membrane three times with 1x TBST for 10 minutes each.

Signal detection

• Prepare an ECL substrate according to the manufacturer’s instructions.
• Incubate the membrane completely with the substrate for 1–5 minutes (adjust the time for more sensitive ECL substrates e.g. SuperSignal West Femto Chemiluminescent Substrate [Pierce]).
• Expose the membrane to autoradiography film in a dark room or read using a chemiluminescence imaging system.

Line up the developed film in the correct orientation to the blot and mark the bands of the MW ladder directly onto the film. It is also advised to add notes such as lane content, film exposure time, and ECL properties.

Immunohistochemistry Technique

Immunohistochemical staining is a valuable tool for detecting specific antigens in tissues. In order to perform the standard staining procedure, first the tissue section has to be deparaffinized and then rehydrated before applying the primary antibody. Enzyme-conjugated secondary antibodies are then applied and the specific staining can be visualized after adding the enzyme-specific substrate. Occasionally, when weak or no staining is observed, an antigen “unmasking” by enzyme digestion, may be required.

The following procedure describes the application of peroxidase or alkaline phosphatase conjugates in the immunohistochemical labeling of formalin-fixed, paraffin-embedded tissue sections.

IHC Protocol Steps

1. Slide Preparation
2. Primary Antibody Reaction
3. Secondary Antibody Reaction
4. Substrate Preparation
5. Development
6. Counterstaining
   

Slide Preparation

Deparaffinization and Rehydration

1. Place the slides in a 56-60 °C oven for 15 min. (Caution: Oven temperature must not exceed 60 °C).
2. Transfer to a xylene bath and perform two changes of xylene for 5 min. each.
3. Shake off excess liquid and rehydrate slides in two changes of fresh absolute ethanol for 3 min. each.
4. Shake off excess liquid and place slides in fresh 90% ethanol for 3 min.
5. Shake off excess liquid and place slides in fresh 80% ethanol for 3 min.
6. Rinse the slides in gently running tap water for 30 seconds (avoid a direct jet which may wash off or loosen the section).
7. Place in PBS wash bath for further rehydration (30 min. at room temperature)

Antigen Retrieval – Unmasking of Antigen

Note: This step is performed only in cases where weak or no staining occurs, or for antigens requiring “unmasking” according to the primary antibody specifications.

There are several possible ways for retrieval depending on the antibody and the antigen:

Enzyme retrieval:

1. Apply 0.1% trypsin in PBS or 0.1% protease in PBS for 2-30 min. at 37 °C. Extending the incubation time may also enhance specific staining. Rinse in PBS for 10 min.

Microwave retrieval:

1. Wash the slides with deionized H2O and place them in a microwave-resistant plastic staining jar containing antigen retrieval solution. Make sure slides are fully covered with solution.
2. Operate the microwave oven for 5 min. on high power (~700 watts). Make sure slides are still covered with retrieval solution or add fresh solution and repeat microwaving.
3. This process can be repeated 2-3 times.
4. Let cool slowly at room temperature for at least 20 min. before proceeding to the next step.

Inactivation of Endogenous Peroxidase

Note: This step is performed only when using peroxidase-conjugated secondary antibodies or ExtrAvidin-Peroxidase.

1. Place the slides on a flat level surface. Do not allow slides to touch each other. Do not allow the sections to dry out at any time.
2. Add enough drops of 3% hydrogen peroxide to cover the whole section.
3. Incubate 5 min. at room temperature.
4. Rinse with PBS from a wash bottle.
5. Place the slide in PBS wash bath for 2 min.

Primary Antibody Reaction

Note:

a. Pre-incubation of the sample with 5% BSA for 10 min. prior to the primary antibody reaction may decrease background staining. For best results with animal tissues, use 5 to 10% normal serum from the same species as the host of the secondary antibody.
b. Optimal dilution and incubation times should be determined for each primary antibody prior to use.

1. Allow the slides to drain, shake off excess fluid with a brisk motion and carefully wipe each slide around the sections.
2. Dilute the primary antibody or negative control reagent to its optimal dilution in diluent. The diluent alone may be used as a negative control. A positive control slide (a tissue known to contain the antigen under study) should also be run.
3. Apply 100 µl primary antibody solution to the appropriate slides, covering the tissue sections.
4. Tilt each slide in two different directions, so the liquid is spread evenly over the slide.
5. Incubate for at least 60 min. at 37 °C in humidified chamber. Longer incubations are advised for low density antigens.
6. Rinse gently with PBS from a wash bottle. Place the slide in a PBS wash bath for 5 min.

Secondary Antibody Reaction

Option 1 – Biotin/ExtrAvidin Detection

1. Allow the slides to drain, shake off excess fluid and carefully wipe the slide as before.
2. Dilute the biotinylated secondary antibody in diluent to its optimal concentration.
3. Apply 100 µl to each slide, covering the tissue sections.
4. Tilt each slide in two different directions.
5. Incubate in a humidity chamber for at least 30 min. at room temperature.
6. Rinse gently with PBS from a wash bottle.
7. Place the slide in a PBS wash bath for 5 min.

ExtrAvidin Reaction

1. Allow the slides to drain. Shake off excess fluid and carefully wipe the slide as before.
2. Dilute ExtrAvidin peroxidase or ExtrAvidin alkaline phosphatase, in diluent to its optimal concentration.
3. Apply 100 µl to all slides; cover the section.
4. Tilt each slide in two different directions.
5. Incubate in humidified chamber for at least 20 min. at room temperature.
6. Rinse gently with PBS from a wash bottle.
7. Place the slide in PBS wash bath for 5 min.

Option 2 – Enzyme-labeled Secondary Antibody

1. Allow the slide to drain. Shake off excess fluid with a brisk motion and carefully wipe the slide as before.
2. Dilute the peroxidase or alkaline phosphatase conjugated secondary antibody in the diluent to its optimal dilution.
3. Apply 100 µl to all slides, covering the tissue sections.
4. Tilt each slide in two different directions.
5. Incubate 30 min. at room temperature or at 37 °C in humidified chamber.
6. Rinse gently with PBS from a wash bottle.
7. Place the slides in a PBS wash bath for 5 min.

Substrate Preparation

It is recommended to prepare the substrate mixture during the final wash step.

Development

1. Allow each slides to drain. Shake off excess fluid and carefully wipe the slide as before.
2. Apply enough drops of freshly prepared substrate mixture to cover the tissue section.
3. Incubate 5-10 min. or until the desired color reaction is observed when monitored with the microscope. Terminate the reaction before background staining appears in the negative controls by rinsing gently with distilled water from a wash bottle.

Counterstaining

Note: When using AEC substrate, do not use alcohol-containing solutions for counter-staining (e.g., Harris’ hematoxylin, acid alcohol), since the AEC stain formed by this method is soluble in organic solvents. The slide must not be dehydrated, brought back to toluene (or xylene), or mounted in toluene-containing mountants.

1. Apply enough Mayer’s hematoxylin to cover the section or place the slide in a bath of Mayer’s hematoxylin.
2. Incubate for 0.5-5 min., depending on strength of the hematoxylin used.
3. Rinse the slide gently with distilled water from a wash bottle.
4. Rinse the slide under gently running tap water for 5 min. (avoid a direct jet which may wash off or loosen the section).
5. Mount the sections using an aqueous mounting medium such as glycerol gelatin. Coverslip may be sealed with clear nail polish.

Flow Cytometry Protocol

This protocol is a recommendation only. Please optimize the procedure since experimental conditions can vary for different samples.

Preparation

1. Harvest and wash the cells and determine the total cell number.
2. Resuspend the cells to approximately 1×106 cells/mL in ice cold PBS. Add the appropriate volume of paraformaldehyde to obtain a final concentration of 4%.
3. Fix for 10 minutes in a 37°C water bath.
4. Wash cells twice with PBS containing 0.5% BSA.
   Note: For extracellular staining with antibodies that do not require permeabilization, proceed to step 8; for intracellular staining, proceed to step 5.

Permeabilization

1. Add ice-cold 90% methanol (approximately 1mL per 1×106 cells) and vortex.
2. Permeabilize for a minimum of 10 minutes on ice.

Immunostaining and Analysis

1. Wash cells twice with PBS containing 0.5%BSA.
2. Resuspend 1×106 cells in 100μL PBS containing 0.5%BSA.
3. Add the primary antibody and incubate for 1 hour at room temperature (RT).
4. Wash cells twice with PBS containing 0.5%BSA.
   Note: If using a fluorescent primary conjugated antibody, skip to step 14.
5. Resuspend cells in fluorochrome conjugated secondary antibody diluted in PBS containing 0.5% BSA.
6. Incubate for 30 minutes at RT.
7. Wash cells twice with PBS containing 0.5% BSA.
8. Resuspend cells in 0.5mL PBS and analyze on flow cytometer.

Direct ELISA Protocol

This protocol is a recommendation only. Please optimize the procedure since experimental conditions can vary for different samples.

Coating Antigen to Plate

1. Coat the wells of a 96 well plate with 100μL of the desired antigen diluted in bicarbonate/carbonate solution. Include a serial dilution of the antigen for analysis.
2. Cover plate with parafilm or plastic adhesive and incubate overnight at 4°C.
3. Remove coating solution and wash plate 2 times with 200μL Phosphate-buffered saline +0.05% Tween20 (PBST). The solutions or washed are removed by flicking the plate over a sink. The remaining drops are removed by patting the plate on a paper towel.

Blocking

1. Block the remaining protein binding sites in the coated wells by adding 200μL of blocking buffer(1% milk/PBS or 1%BSA/PBS).
2. Cover the plate and incubate for 2 hours at room temperature (RT).
3. Wash plate 2 times with 200μL PBST.

Antibody Incubation

1. Add 100μL of antibody diluted in blocking buffer.
2. Cover the plate and incubate at RT for 2 hours.
3. Wash plate 2 times with 200μL PBST.

Detection

1. Add 100μL of the substrate solution per well.
2. After sufficient color development add 100μL of stop solution to the wells.
3. Read the absorbance of each well with a plate reader.

Analysis

1. Prepare a standard curve from the data produced from the serial dilutions with concentration on the X-axis vs. absorbance on the Y-axis. Interpolate the concentration of the sample from this standard curve.

Indirect ELISA Protocol

Coating Antigen to Plate

1. Coat the wells of a 96 well plate with 100μL of the desired antigen diluted in bicarbonate/carbonate solution. Include a serial dilution of the antigen for analysis.
2. Cover plate with parafilm or plastic adhesive and incubate overnight at 4°C.
3. Remove coating solution and wash plate 2 times with 200μL Phosphate-buffered saline +0.05% Tween20 (PBST). The solutions or washed are removed by flicking the plate over a sink. The remaining drops are removed by patting the plate on a paper towel.

Blocking

1. Block the remaining protein binding sites in the coated wells by adding 200μL of blocking buffer(1% milk/PBS or 1%BSA/PBS).
2. Cover the plate and incubate for 2 hours at room temperature (RT).
3. Wash plate 2 times with 200μL PBST.

Antibody Incubation

1. Add 100uL of antibody diluted in blocking buffer.
2. Cover the plate and incubate for 2 hours at RT.
3. Wash plate 2 times with 200μL PBST.
4. Add 100uL of conjugated secondary antibody diluted in blocking buffer.
5. Cover the plate and incubate for 2 hours at room temperature.
6. Wash plate 2 times with 200μL PBST.

Detection

1. Add 100uL of the substrate solution per well.
2. After sufficient color development add 100μL of stop solution to the wells.
3. Read the absorbance of each well with a plate reader.

Anaylsis

1. Prepare a standard curve from the data produced from the serial dilutions with concentration on the X-axis vs. absorbance on the Y-axis. Interpolate the concentration of the sample from this standard curve.

Sandwich ELISA Protocol

Coating with Capture Antibody

1. Coat the wells of a 96-well plate with 100μL of the capture antibody diluted in bicarbonate/carbonate solution.
2. Cover the plate with parafilm or plastic adhesive and incubate overnight at 4°C.
3. Remove the coating solution and wash plate 2 times with 200μL Phosphate-buffered saline +0.05% Tween20 (PBST). The solutions and washes are removed by flicking the plate over a sink. The remaining drops are removed by patting the plate on a paper towel or by aspiration.

Blocking

1. Block the remaining protein-binding sites in the coated wells by adding 200μL blocking buffer per well.
2. Cover the plate and incubate for 2 hours at room temperature (RT).
3. Wash the plate 2 times with 200μL PBST.

Adding Samples

1. Add 100uL of appropriately diluted samples and standards.
   Note: For accurate quantitative results, always compare signal of unknown samples against those of a standard curve. Standards (in duplicate and triplicate) and a blank must be run with each plate for analysis and to ensure accuracy.
2. Cover the plate and incubate for 2 hours at room temperature.
3. Wash the plate 2 times with 200μL PBST.

Incubation with Detection Antibody

1. Add 100uL of diluted detection antibody to each well.
   Note: Be sure to check that the detection antibody recognizes a different epitope on the target protein to the capture antibody. This prevents interference with antibody binding.
2. Cover the plate and incubate for 2 hours at room temperature.
3. Wash the plate 4 times with PBS.
4. Add 100μL of conjugated secondary antibody, diluted in blocking buffer immediately before use.
5. Cover the plate and incubate for 2 hours at room temperature.
6. Wash the plate 4 times with PBS.

Detection

1. Add 100μL of the substrate solution per well.
2. After sufficient color development add 100μL of stop solution to the wells.
3. Read the absorbance of each well with a plate reader.
4. Prepare a standard curve from the data produced from the serial dilutions with concentration on the X-axis vs. absorbance on the Y-axis. Interpolate the concentration of the sample from this standard curve.

Immunoprecipitation Protocol

This protocol is a recommendation only. Please optimize the procedure since experimental conditions can vary for different samples.

Sample Preparation

Non-Denaturing:

1. Place cell culture dish on ice and wash the cells with ice cold Phosphate-buffered saline (PBS).
2. Drain PBS and add ice cold lysis buffer.
3. Scrape cells off the dish using a cell scraper and gently transfer the cell suspension into a microcentrifuge tube.
4. Maintain agitation for 30 minutes at 4°C.
5. Centrifuge in a micro centrifuge at 4°C. The time and force of centrifugation may vary depending on the cell type.
6. Gently remove the tubes from the centrifuge and place them on ice.
7. Aspirate the supernatant ad place in a fresh tube on ice.

Denaturing:

1. Add 100uL denaturing lysis buffer to the cells.
2. Mix well by vortexing vigorously for 2 to 3 seconds.
3. Transfer the cell suspension to a microcentrifuge tube.
4. Heat samples to 95°C for 5 minutes to denature.
5. Dilute the suspension with 0.9mL of non-denaturing lysis buffer. Mix gently.
6. Fragment the DNA by passing the lysed suspension 5-10 times through a needle attached to a syringe.
7. Incubate on ice for 5 minutes.

Tissue lysates:

1. Dissect the tissue quickly with clean tools. If possible, do so on ice to prevent degradation.
2. Place tissue in microcentrifuge tubes and snap freeze by immersing in liquid nitrogen.
3. Add 300uL of lysis buffer for approximately 5mg of tissue and homogenize with an electric homogenizer.
4. Rinse twice with another 300uL of lysis buffer per rinse and maintain constant agitation for 2 hours at 4°C.
   Note: If denaturing is required, follow steps 2-5 from the denaturing protocol above.
5. Centrifuge for 20 minutes at 12000rpm at 4°C in a microcentrifuge. Aspirate the supernatant and place in a fresh tube kept on ice.

Pre-clearing the Lysates

1. Add 50uL of normal serum to 1 mL of lysate and incubate for 1 hour on ice.
2. Add 100uL of bead slurry to the lysate and incubate for 30 minutes at 4°C.
3. Spin in microcentrifuge at 14,000 x g for 10 minutes at 4°C.
4. Discard bead pellet and keep supernatant for immunoprecipitation.

Immunoprecipitation with Antibody in Solution

1. While on ice, add 10-50ug cell lysates to a microcentrifuge tube plus the recommended amount of antibody.
2. Incubate the sample with the antibody for a few hours at 4°C with gentle agitation.
   Note: The length of incubation depends on the amount of protein and the affinity properties of the antibody.
3. Prepare the Sepharose beads.
4. Mix the slurry well and add 70-100uL of the beads to each of the samples, while on ice.
5. Incubate the beads mixture for 4 hours at 4°C with gentle agitation.
6. Centrifuge tube after incubation and wash 3 times with lysis buffer.
7. Add 25-50 uL 2x loading buffer and boil for 5 minutes at 100°C.
8. Centrifuge and transfer the supernatant to a new tube for Western blotting.