Western Blotting Guide

What is Western Blotting?

Western blotting is a technique used to determine the presence or absence of selected proteins in a sample. Western blot is advantageous over other antibody based detection assays such as ELISA as cross-reactivity with non-target proteins can be differentiated from the target antigen based on molecular weight.

First the proteins are separated on a basis of size by gel electrophoresis. Following this the protein is transferred to a membrane usually nitrocellulose or PVDF, through the use of an electrical current. The membrane is then stained with antibodies specific for the protein of interest, enabling the acquisition of qualitative or semi-quantitative information about the protein.

1. Sample preparation – Protein Extraction

The first step of a western blot protocol is protein extraction from cells or tissue. The protein of interest must be solubilized in order to migrate through the separating gel.

The choice of lysis buffer used will depend on the yield of protein required and the subcellular localisation of the protein. Lysis buffer containing sodium dodecyl sulfate (SDS) and other ionic detergents is considered to give the highest protein yield and is the most damaging to the sample. It should also be noted that the lysis buffer used will affect antibody choice further on in the protocol with regards to the protein form it recognizes, either native or denatured.

Lysis buffers containing SDS have a denaturing effect on protein, whilst buffers without detergent or mild non-ionic detergents such as NP-40 and Triton X-100 should be used when the antibody will only recognise protein in its native structure. Information about the protein form your antibody recognises can be found on the data sheet supplied by the manufacturer.

It is important to note that when preservation of protein-protein interactions is required a buffer without ionic and non-ionic detergents should be used. This may be achieved by mechanical shearing.

Protein Localization Recommended Buffer
Cytoplasmic (cytoskeletal bound)
Cytoplasmic (soluble)
Membrane Bound
Whole Cell

1.1 RIPA Buffer

RIPA (Radio Immuno Precipitation Assay) buffer is used to lyse and extract protein from cultured cells. RIPA buffer is an ideal cell lysis reagent for whole cell extracts and membrane-bound proteins. As RIPA buffer will disrupt protein-protein interactions it may not be ideal for immunoprecipitation assays.

1.1.2 Sample RIPA Buffer Recipe

50mM Tris HCL pH 7.4, 50 mM NaCl, 2mM EDTA, 0.1% SDS. Plus freshly added proteinase Inhibitors (Apoprotein, Leupeptin, DTT and PMSF)

1.2 NP-40 Buffer

NP-40 buffer is widely used for the extraction of cytoplasmic, membrane-bound and whole cell extracts. NP-40 is considered a weaker buffer to RIPA buffer.

1.2.1 Sample NP-40 Buffer Recipe.

150 mM sodium chloride, 1.0% NP-40 (Triton X-100 can be substituted for NP-40), 50 mM Tris, pH 8.0

1.3 Tris-HCL Buffer

20 mM Tris-HCl, pH 7.5

1.4 Tris-Triton Buffer

  • 10 mM Tris, pH 7.4
  • 100 mM NaCl
  • 1 mM EDTA
  • 1 mM EGTA
  • 1% Triton X-100
  • 10% glycerol
  • 0.1% SDS
  • 0.5% deoxycholate

*All buffers can be stored at 4C for several weeks and at -20C for up to a year.

1.5 Protease and Phosphatase Inhibitors

Once cell lysis occurs so too does protein degradation. To prevent proteolysis and dephosphorylation the samples must be kept on ice at all times. Proteinase and phosphatase inhibitors are added to the lysis buffer to slow this process down. These inhibitors must be added freshly each time to the lysis buffer.

Inhibitor Target
Chymotrypsin, Plasmin, Trypsin
Metalloproteases requiring Mg++ and Mn++
Metalloproteases requiring Ca++
Na Fluoride
Serine/Threonine Phosphatases
Na Orthovandate
Tyrosine phosphatases
Pepstatin A
Aspartic proteases
Serine, Cysteine proteases

1.6 Sample protocol- Preparation of cell lysate from cell culture using RIPA Buffer

Steps Procedure


Wash cells with ice cold PBS


Aspirate PBS


Add ice cold RIPA Buffer (~1ml per 107 cells)


Scrape adherent cells off the plate using your sterile pipette tip


The centrifugation force and time can vary depending on cell type.


Remove from centrifuge and store on ice.


Aspirate the supernatant into a new tube and keep on ice, discard the pellet.


Determine protein concentration using a Bradford assay, a Lowry assay or a bicinchoninic acid (BCA). BSA can be used a standard


Once the protein concentration has been determined the samples can be frozen at -20 °C / -80 °C or prepared for loading.

1.7 Bradford Assay for protein determination

Step Procedure


Aliquot (2 ml) of each protein sample was into separate wells of a 96 well plate.


Add Bradford reagent (100 ml) to each sample. Ensure that no bubbles are introduced.


Gently shake at room temperature for 5 min.


To calculate the protein concentration in each sample read the absorbance off a BSA standard curve, constructed as follows: prepare serial dilutions of BSA between 2 mg/ ml and 15 mg/ml and add to 100 ml of Bradford reagent in a 96 well plate.


Measure absorbance at 595 nm, normalise to a reference measurement at 450 nm against the blank (2 ml lysis buffer, 100 ml dye reagent) on a Pro-Max5 microplate reader.


The protein sample concentration can be calculated against the standard curve.

1.8 Preparation of samples for loading into gels

In some cases, the antibody used will recognise the protein in native state, as the selected epitope may exist on the surface of the folded structure. When this is the case it is not necessary to denature the sample and therefore SDS should be left out and the sample. Additionally, reducing agents such as ß-mercaptoethanol and DTT must be left out of the loading buffer and migration buffer.

Certain proteins will require denaturation for the antibody to work effectively. Heat denaturation will unfold the protein and enable the antibody to bind its corresponding epitope located within the 3D conformation of the protein. Protein denaturation can be achieved using a loading buffer containing a denaturing agent e.g SDS which is heated at 95-100°C for 5 minutes

The standard loading buffer for western blot samples is 2x Lameli Buffer. Lameli Buffer contains beta-2-mercaptoethanol or dithiothreitol (DTT) which act to reduce disulphide bonds before they adopt the random-coil configuration and in turn denatures the protein. Lameli buffer also has a SDS component which provides the negative charge necessary for gel electrophoresis, in addition to glycerol to make the sample denser.

To enable visualization of the migration of proteins it is common to include small anionic dye molecule (e.g., bromophenol blue) in the loading buffer. As the dye is anionic and small it will migrate the fastest of any component in the mixture to be separated and provide a migration front to monitor the separation progress.

1.8.1 Lameli Buffer Recipe

4% SDS, 10% 2-mercaptoehtanol., 20% glycerol, 0.004% bromophenol blue, 0.125 M Tris HCl. Check the pH and bring it to pH 6.8.

2. Electrophoresis

Once the sample has been lysed, the protein concentration determined and loading buffer added the sample can now be separated by gel electrophoresis.

There are 2 parts to an SDS-PAGE gel for a western blot. The resolving and stacking gel. The bottom and larger portion of the gel is known as the resolving gel – this is poured first. Once the resolving gel has set (allow up an hour) the stacking gel can be added. The stacking gel is poured on top of the resolving gel – the combs are placed into this gel. The basic chemistry of these two gels is identical they only differ in their acrylamide concentration and pH.

The percentage acrylamide used in the gel depends on the size of the protein of interest and size of pore needed in the gel. Generally, for small proteins a high percentage gel should be used, for larger proteins a low percentage gel should be used. It should be noted that as the amount of acrylamide used increases the pore size decreases. The below table outlines the recommended percentage gels based on linear separation range.

Acrylamide Concentration % Linear Range of Separation (KD)

Gels can be bought; however many research groups prefer to make their own. Below is a sample recipe for 10% stacking gel and resolving gel.

2.1 10% Resolving Gel Recipe

Reagent Volume


5.9 ml
30% Acrylamide-Bis
5 ml
1.5M Tris pH 8.8
3.8 ml
10% SDS
150 µl
10% APS
150 µl
6 µl

2.2 Western Blot Stacking Gel Recipe

Reagent Volume
2.7 ml
670 µl
Tris pH 6.8
500 µl
10% SDS
40 µl
10% APS
40 µl
3 µl

3. Controls and Molecular Weight Markers

As with all experiments a positive and negative control should be used to determine that the assay is accurate, sensitive and efficient. Loading controls such also be used to ensure that the gel has been evenly loaded.

Molecular weight markers

Using a range of molecular weight markers will enable the determination of the protein size (see below) and enable the electrophoretic run to be monitored.

Sample Type Protein MW (kDa)

Whole cell / cytoplasmic proteins

beta actin
alpha actin
beta tubulin
alpha tubulin

High Molecular Weight (HMW)



cytochrome C oxidase

Nuclear proteins

lamin B1
TATA binding protein TBP
histone H1


histone H3


Plant tissue





Serum sample


Muscle sample

SDHA [7]

Yeast sample

Phosphoglycerate kinase


4. Sample Protocol for Loading Samples and Running the Gel

Step Procedure


Separate proteins by SDS-PAGE on an acrylamide gel. The percentage of acrylamide used will depend on the molecular size of your protein of interest. In this sample protocol a 10% gel is described. In the table below you will find a guide to acrylamide concentration and best practice separation ranges for proteins.


Poor the resolving gel between the plates and let the gel set for 1 hr.


Level the gel with isopronol. For this take 300 uL of Isopropanol and poor carefully across the top of the gel.


Once your gel has set you can remove the Isopropanol by puring a gentle flow of water over the gel under the tap.


Add the stacking gel and carefully place your comb into the gel, avoiding air bubbles. Allow gel to set for 1 hr.

Step Procedure


Poor the resolving gel between the plates and let the gel set for 1 hr.


Level the gel with isopronol. For this take 300 uL of Isopropanol and poor carefully across the top of the gel.


Once your gel has set you can remove the Isopropanol by puring a gentle flow of water over the gel under the tap.


Add the stacking gel and carefully place your comb into the gel, avoiding air bubbles. Allow gel to set for 1 hr.


Place the plate in the gel electrophoresis rig and add 400ml of 1x SDS-PAGE running buffer (15.1 g TRIZMA, 94 g glycine,50 ml of 10% w/v SDS,dH20 to 1 litre), first to the inside of the chamber and let over flow out to ensure no leakage.


Remove the combs and wash out the wells with running buffer if necessary.


Add 8 µl of unstained molecular weight ladder and 20 µl of the protein samples of interest to the selected well.


Run the gels at 25mA per gel for 1 hr 15 min

5. Western Blot Transfer

Following electrophoresis, the protein must be transferred from the gel to a membrane, either nitrocellulose or PVDF. The most widely used transfer method is electroelution or electrophoretic transfer. This method relies on the hydrophobicity and electrostatic charges between the protein and the PVDF or nitrocellulose membrane, determining the efficacy of protein transfer.

The protein-containing gel is placed in direct contact with the transfer membrane and sandwiched between 2 electrodes submerged in a conducting solution. When an electric field is applied the proteins migrate from the gel and attach to the membrane. The membrane is now a copy of the protein pattern observed on the polyacrylamide gel.

*Note- Its best to always probe for your weakest antibody first, as handling and stripping can unstick your proteins of interest.

5.1 Transfer Buffer recipe

5.8g Trizma base, 2.9g glycine, 0.37g SDS powder, 200ml methanol dH20 to 1L

6. Blocking

Before the antibody is added it is necessary to block the membrane to prevent any nonspecific binding of the detection antibody. Blocking the membrane improves the sensitivity and signal-to-noise ratio of your assay. Milk and BSA are the most commonly used blocking agents.

The blocking agent used will depend on the antibody as some antibodies have been shown to non-specifically to bind to milk. When probing with phosphorylation sensitive antibodies it is recommended to use BSA.

It is important to note that if your primary antibody was made against a horse, cow, goat or donkey that you cow proteins BSA or milk can’t be used to block the membrane due to the potential for cross reaction or IgG contamination.

6.1 Blocking buffer recipe

5% marvel in TBS-Tween (1X TBS with 0.1% v/v tween-20)

7. Wash Buffers

Wash steps in a western blot are necessary to remove any unbound reagents and reduce background. Failure to wash properly can result in a high background noise, too much washing can elute the antigen from the blot.

The amount of detergent used in your wash buffer can vary depending of your assay with concentrations ranging from 0.05 to 0.5% for detergents like Tween 20. It is recommended to make fresh stocks of detergents e.g. Tween 20 as microbial growth can result in high background noise. Furthermore, as detergents contain peroxides it is essential to use high-purity detergents.

7.1 Wash Buffer – TBS-T Recipe

  • 20 mM Tris, pH 7.5
  • 150 mM NaCl
  • 01% Tween 20

8. Primary Antibody Incubation

Once the membrane has been blocked to prevent non-specific binding it can now be probed with the primary antibody. A good primary antibody will recognise a specific protein or epitope on a group of proteins. The antibody used will depend on the protein under investigation.

When choosing a primary antibody a number of considerations must be taken into account, is the antibody species specific as epitope can vary between species, the antibody/protein interaction may be sensitive to denaturing conditions and post-translational modifications and finally not all primary antibodies are suitable for western blotting and may need verification.

Following incubation with primary antibody the membrane should be washed x5 times for 5 minutes with wash buffer

9. Secondary Antibody Incubation

In general, as the primary antibody is not detectable a labelled secondary antibody must be used which binds the primary antibody and enables detection of the target antigen. The choice of secondary antibody used can depend on the species the primary antibody was raised in or the tag linked to the primary antibody e.g biotin, histidine. For instance, if the primary antibody is an unmodified rabbit monoclonal antibody the secondary antibody used must be an anti-mouse secondary antibody from a non-mouse host.

As mentioned above secondary antibodies are conjugated to different developing molecules, for film detection and chemiluminescence HRP or AP is commonly used whilst for laser capture a fluorophore is necessary.

10. Antibody Dilution

The dilution at which the antibody should be used at is commonly recommended by manufacturers. However, the optimal dilution of antibody can vary depending on the specifications of the assay. More strongly expressed proteins and assays of high sensitivity will require less antibody in comparison to more weakly expressed proteins and less sensitive assays. Using less antibody decrease background noise and increase specificity. Antibody dilutions are normally made in wash buffer.

11. Blot Development

The type of label on the secondary antibody will determine the development system used. For enzymatically labelled antibodies e.g. HRP and AP a chemiluminescence method is needed. If a conjugated fluorophore was used a fluorescent scanning device can be used instantaneously.

11.1 Chemiluminescence

HRP conjugated antibodies are the most widely used and considered superior to AP conjugates because of the specific activities of both enzyme and antibody due to the smaller size of the HRP enzyme and its compatibility with conjugation reactions. Furthermore, HRP substrates are widely available in addition to offering a high activity rate and stability.

AP although possessing a linear reaction rate the increased reaction time required can often result in a high background signal and low signal-to-noise ratio.

The signal generated from HRP/AP conjugates is transient and only persists as long as the enzyme-substrate reaction is occurring. A well optimised assay of antibody dilution and sufficient substrate can produce light for 1 to 24hrs, allowing for documentation either with X-ray film or digital imaging equipment. X-ray film can be used to obtain semi-quantitative data but digital imaging allows for a detection of broad dynamic range and enables the retrieval of quantitative data for western blots. In chemiluminescent blotting the signal obtained is variable and generally regarded as semi-quantitative.

11.2 Fluorescent Detection

Fluorophore conjugated antibodies require fewer steps as no substrate is used making the protocol much shorter. Specialised equipment is however required to detect the signal requiring an excitation light source. The development of infrared, near-infrared and quantum dots has increased the sensitivity of fluorescent probed for western blot analysis.

An advantage of using fluorescent detection over chemiluminescent detection is the ability to multiplex using more than one fluorophore. Florescent blotting also enables quantitative and consistent results across repeat experiments. Disadvantages of fluorescent detection can include a low signal to noise ratio as a result of auto- fluorescence and a failure to amplify lowly expressed proteins when using a charged-couple device camera.

11.3 Differences between Chemiluminescent Detection and Fluorescent Detection

Chemiluminescent detection Fluorescent Detection


Secondary antibody labelled with enzyme (e.g. HRP or AP)
Secondary antibody labelled with fluorophore

Method of Detection

X-Ray film exposure/ Digital Imaging
Laser Scanning Imager

Ability to multiplex




Strength of signal

Weeks to Months

Linear Dynamic Range

15-fold (X-Ray Detection), 3,000-4,000- fold (Digital Imager)

Quantitative Measurement

Enzymatic signal is variable and only semi-quantitative
Fluorescent signal is static and quantitative


No substrate needed

12. Stripping and Re-probing of Blots

The removal of primary and secondary antibodies from a western blot membrane is known as stripping. Stripping enables the investigator to look at more than 1 protein on the blot e.g the protein of interest and a loading control. Probing can also be used for multiple targets and is advantageous as it saves time and sample.

Stripping works best on a PVDF membrane as it has greater capabilities to keep the protein attached and not elute. Chemiluminescent reagents such as ECL are recommended for stripping as they don’t stain the membrane which can interfere with the detection of targets of similar molecular weights. How many times a membrane can be stripped and stained depends on how optimised the stripping protocol is.

Following stripping it is important to thoroughly was the membrane with buffer and block it before primary antibody incubation.

There 2 different strengths of stripping buffer which can be used mild or harsh.

12.1 Mild Stripping Buffer

A mild stripping buffer uses a low pH glycine solution to dissociate bound antibodies. The low pH acts to remove bound antibodies via structural alteration and deactivation of the active site.

12.2 Mild Stripping Buffer recipe

15 g glycine
1 g SDS
10 mL Tween 20
Dissolve in 800 mL distilled water
Adjust pH to 2.2
Bring volume up to 1 L with distilled water

12.3 Harsh Stripping Buffer

A harsh stripping is primarily used on blots with a high signal strength. Harsh stripping buffers also rely on a low pH to alter antibody structure and release the target protein. In harsh buffers this activity is achieved through the use of a neutral Tris-HCl solution containing reducing agents such as beta-mercaptoethanol and SDS. This solution is heated with the blot at 50–80°C for up to 45 minutes with agitation. It should be noted that the activities of SDS and beta-mercaptoethanol can’t be reversed and the stripped antibody recovered. Furthermore, thorough washing is required following the use of a harsh buffer to prevent denaturation of the re-probing antibody.

12.4 Harsh Stripping Buffer Recipe

62.5 mM Tris-HCL,pH 7.8, 100 mM -mercaptoethanol, 2% (w/v) SDS

12.5 Sample Striping protocol

Steps Procedure


Incubate the PVDF Membrane with stripping buffer (62.5 mM Tris-HCL,pH 7.8, 100 mM -mercaptoethanol, 2% (w/v) SDS) for 30 min at 50 °C.


Wash the membrane 2 X 5 min, 1 X 10 min, 2 X 5 min at room temperature with agitation on a platform rocker.


Block by incubation with gentle agitation in blocking solution for 1 h at room temperature.


Washed 2 X 5 min, followed by incubation with the appropriate primary and secondary antibody.

13. Protein Gel Visualisation

The separation of protein by gel electrophoresis can be visualised in 2 ways: Coomassie staining or copper staining. The choice of staining used depends largely on the downstream applications of the protein.

13.1 Coomassie Staining

Coomassie staining is used to determine whether the protein has migrated uniformly and evenly. A coomassie stain should only be used on gel if the aim is not to transfer but to observe the results of SDS-Page separation as coomassie staining not reversible.

13.2 Coomassie Staining Sample Protocol

Steps Procedure


Following gel electrophoresis treat the gel with 40% distilled water, 10% acetic acid, and 50% methanol, this will the proteins to precipitate.


Place the gel in the same solution as used in step 1 but with the addition of 0.25% by weight Coomassie blue stain.


Incubate for 4-24 hrs at room temperature on a shaker.


Transfer the gel to a mixture of 67.5% distilled water, 7.5% acetic acid, and 25% methanol on shaker to rinse.


Replace with fresh rinse buffer once the excess dye has been removed.


The stain will not bind to the acrylamide, and will wash out (leaving a clear gel).


The stain binds strongly to the proteins in the gel and give off a blue colour.

13.3 Copper Staining

Copper staining should be used if you wish to transfer the protein following visualization. The copper staining is said to be faster and more sensitive than coomassie staining.

13.4 Copper Staining Sample Protocol

Steps Procedure


Briefly rinse the gel in ddh20 following electrophoresis


Transfer to a solution of 3 M CuCl2for 5–15 min


Wash the gel in ddh20


View against a dark field background


The protein can be noted as clear zone in a translucent blue background


Following visualisation gels can be de-stained by washing in 0.1–0.25 M Tris/0.25 M EDTA pH 8.0


Place the gel in transfer buffer and proceed with the transfer

14. Protein Membrane Visualisation

The efficiency and success of the transfer can be assessed by staining the PVDF or nitrocellulose membrane with Ponceau Red. Staining with ponceau red is easily reversed with washing and therefore enable subsequent antibody probing.

14.1 Ponceau Red Sample Protocol

Steps Procedure


Dilute Ponceau stock 1:100


Incubate on an agitator for 5 min


Wash with ddh20 until the water is clear and the protein bands visible


Destain the membrane by washing with TBST.


If using a PVDF membrane , re-activate the membrane with methanol then wash again in TBST.