Western blotting or Immunoblotting is a technique which enable to study the distribution and behaviour of proteins in extracts prepared from tissues or cells, based on detection using specific antibodies (Abs). Electroblotting is a technique used to immobilise proteins or nucleic acid separation on a solid membrane support.
How does it work?
Western blotting identifies with specific Abs proteins that have been previously separated from one another according to their size (molecular weight – 1D SDS-PAGE electrophoresis) or/and isoelectric point (2D SDS-PAGE electrophoresis) by gel electrophoresis.
This application notes will give an introduction of the entire process and steps of producing a Western blot. This technique involves several steps which will vary according to the specific starting material, protein location, solubility etc. This application notes will give a general outline of the steps needed to design a Western blot protocol from the starting material to the developing of the blot, however specific steps will vary according to specific needs, thus these are only examples.
To prepare a sample for running on a gel, cells and tissues must be lysed to release the protein of interest. Lysis can be performed through mechanical (i.e. sonication) or enzymatic methods. This will also solubilise the protein allowing them to migrate individually and separate through a gel by electrophoresis. There are a variety of recipes for lysis buffer which differ according to specific proteins and antibodies. Most of the lysis buffer contains denaturing and reducing agents such as sodium dodecyl sulphate (SDS) and other ionic detergents. Most antibodies will recognise proteins in their denatured form, however, it is important to notice that some antibody (it will be specified on the data sheet) will recognise proteins only on their naive form, thus denaturing detergents should be avoided in this case (mild non-ionic detergents such as NP-40 and Triton X-100 can be used if necessary).
Examples of buffers:
|NP-40 or RIPA or Tris-HCl
|Cytoplasmic (cytoskeletal bound)
|NP-40 or RIPA or Tris-Triton
|RIPA or nuclear fraction protocol*
|RIPA or mitochondria fraction protocol *
*Proteins that can be found on sub-cellular location can be enriched in the lysate of the sub-cellular fraction using specific protocols. This can be useful when trying to obtain a signal from a weakly-expressed protein.
Protease and Phosphatase inhibitor
Following lysis proteolysis, phosphorylation and denaturation begins, therefore it is fundamental, in order to slow down these events, to keep the samples at 4oC at all the time and freshly add to the lysis buffers appropriate inhibitors.
Examples of Inhibitors:
|Final concentration in lysis buffer
|Stock (store at -20oC)
|Trypsin, chymotrypsin, plasmin
|Dilute in H2O, 10mg/ml
|Dilute in H2O
|Dilute in methanol, 1mM
|Serine, cysteine, proteases
|Dilute in ethanol
|Metalloproteases that require Mg2+ and Mn2+
|Dilute in dH2O, 0.5M, adjust to pH8
|Metalloproteases that require Ca2+
|Dilute in dH2O, 0.5M, adjust to pH8
|Dilute in water
|Dilute in water
Preparation of lysate
Specific protocols should be used according to the starting material (i.e. tissues, cell culture, bacteria) and the protein of interest, however all the steps should be performed at 4oC. Once the required proteins fractions are obtained they should be stored at -20oC for several weeks or -80oC for longer period.
Proteins can be quantified using several methods. The most common ones are Bradford assay, Lowry assay or BCA assay. Usually the protein standard frequently used to determine the standard curve is BSA (bovine serum albumin).
Alternatively, the proteins concentration can be directly measure using a spectrophotometer at absorbance of 280nm. This method is very fast and don’t require any standards, since the relationship between absorbance and protein concentration is linear. However, they can produce considerable error in protein concentration because non-protein component that absorb the ultraviolet light will interfere, thus it is not recommended when very precise quantification is needed.
Preparation of the sample for loading into gel
Denatured, reduced samples
Antibodies usually recognise a small portion of the specific protein and this domain may reside within the 3D conformation of the protein. Therefore, in order to allow the Abs to access it, the protein must be unfolded (denatured).
Generally, to denature the proteins a loading buffer (Laemmli buffer) containing the anionic denaturing detergent (SDS) is added to the sample.
Laemmli buffer contains the following chemicals and these are their function:
SDS denatures the protein by binding to their polypeptide backbone, conferring a negative charge to the polypeptide which is proportional to its length. Therefore, the migration of the proteins in denaturing SDS-PAGE is determined by molecular weight.
The addition of 2-mercaptoethanol or dithiothreitol (DTT) is important in order to reduce the disulphide bridges in proteins before they adopt the random-coil configuration, which is necessary for separation by size.
Glycerol is added to increase the density of the sample and hence to maintain the sample at the bottom of the well after loading.
Bromophenol blue is an anionic dye commonly included to enable visualisation of the migration of the protein. Since the dye is very small it will migrate faster than any component in the mixture to be separated and provide indication and monitoring of the separation progress. Generally, the run should be stopped when the dye reaches the bottom of the gel.
The mixture is then boiled at 95-100oC for 5 minutes and then loaded onto the gel.
If the samples are stored for future re-use they should not be re-boil again, they can be loaded directly.
Native and non-reduced sample
Some antibody may recognise an epitope of non-contiguous amino acids. Although the amino acids of the epitope are separated in the primary sequence, they are closed to each other in the folded three-dimension structure of the protein. In this case the antibody will only recognise the epitope as it exists on the surface of the folded structure.
In most of these circumstances to run a Western blot in non-denaturing conditions, it simply means leaving the SDS out of the sample and migration buffer and not heating the sample.
Sometime certain antibodies only recognise protein in its non-reduced form, thus, in this case the reducing agents 2-mercaptoethenol or DTT must be left out of the loading buffer and migration buffer.
Electrophoresis can be one dimensional (according to proteins molecular weight) or two dimensional according to protein isoelectric point and molecular weight). 1D dimensional electrophoresis is usually used for most routine protein separation. 2D dimensional separation when it is necessary to resolve all the protein present in the cell or maybe if two proteins of interest have the same molecular weight.
In this note we will describe the technique for 1D SDS-PAGE. For 2-D SDS-PAGE follow manufacturer instructions.
Preparation of 1D PAGE gels
Polyacrylamide gels are formed from the polymerisation of acrylamide and N,N-methylenebis-acrylamide (Bis). Bis is the cross-linking agent for the gel. The polymerisation begins when ammonium persulfate and TEMED are added. The gels are neutral, hydrophilic tree-dimensional networks of long hydrocarbons cross-linked by methylene groups.
The separation of the proteins depends by the relative size of the pores formed within the gel. The pore size is dependent on the total amount of acrylamide present and the amount of cross-linker. As the amount of acrylamide increases, the pore size decreases. Therefore, to separate high molecular weight proteins is better to use low percentage gels, such as 8%, whereas for separation of low molecular weight proteins at 15% gel may be better.
A positive control lysate is used to demonstrate that the Western blot was performed correctly and that the antibody recognises the target protein, which may not be present in the experimental sample.
Molecular weight marker
Molecular weight markers will enable determination of the size of the target protein by comparison. Pre-stained molecular markers are also useful to monitor the progress of proteins separation during electrophoresis and to give an indication on whether or not the transfer onto the membrane (blot) was successful.
Loading samples and running the gel
To load the sample into the well, special gel loading tips or a micro-syringe can be used. N.B. Take care not to poke the well bottom with the tip as this will create distorted bands. Never overfill the well, which could lead to poor data, if samples spill into adjacent well.
Generally, 20-40 mg of total proteins is loaded per mini-gel well.
The gel will be submerged in 1x Tris-glycine Running buffer (other buffers can be used according to the specific needs) and the electrophoresis will proceed until the bromophenol blue tracking dye reaches the bottom of the gel. Running time will vary according to the percentage of the gel and voltage used. Mini-gel can be run at current up to 30mA/gel.
Once the run is finished the proteins will slowly start to elute from the gel, therefore do not store the gel and proceed immediately to transfer.
Use of loading controls
In order to check that the lane have been evenly loaded with sample, loading controls should be used, especially when a comparison must be made between the expression level of a protein in different sample. They are also useful to check whether that transfer was even throughout the whole gel. Moreover, they can be used to quantify the protein amount in each lane, if even load or transfer did not occur. Loading control must be present for publication-quality work.
Below are some examples of loading controls used in mammalian cells.
|Molecular weight (KDa)
|Whole cell/ cytoplasmic
|Not suitable for muscle sample. Changes in cell-growth conditions and interactions with extracellular matrix components may alter actin protein synthesis (Farmer at al. 1983)
|Whole cell/ cytoplasmic
|Some physiological factors, such as hypoxia and diabetes, increase GAPDH expression in certain cell type
|Whole cell/ cytoplasmic
|Tubulin expression may vary according to resistance to antimicrobial and antimitotic drugs (Sangrajrang et al. 1998; Prasad et al. 2000)
|Many proteins run at the same size as COXIV
|Not suitable for sample where the nuclear envelope is removed
|TATA binding protein TBP
|Not suitable for sample where DNA is removed
N.B. Always check publications for accepted loading controls.
Transfer of protein (Western Blotting) and staining
Visualisation of protein on gels
N.B. This step can be usually omitted.
Visualisation of the gel can give an indication if the proteins have migrated evenly. If you plan to transfer the separated protein to a membrane, use Copper stain. Coomassie stain is not reversible (NEVER USE IF YOU PLAN TO BLOT THE GEL).
Coomassie stain can be used if you just want to visualise the protein on the gel or half of the samples on the gel for later comparison with the blotted samples loaded as mirror image.
As already mentioned as soon as the run is finished the proteins will start eluting from the gel. Addition of the Coomassie stain will causes most of the proteins to precipitate preventing their diffusion. The dye will enable their visualisation. The gels can be stained for 4hrs or overnight at room temperature on a shaker. The gel will be then transferred into the de-stain solution (replace with fresh solution as required) which will rinse the excess dye off the gel and allow visualisation of blue protein bands. Once reached the desired look of the protein bands, the gel can be washed several times in distilled water and stored at 4oC for several weeks, alternatively it can be dried and kept for long term storage.
Briefly rinse the gel in distilled water (30 seconds) and transfer it in Copper stain for 5 to 15 minutes. Wash the gel briefly in de-ionised water and view the gel against a dark-field background. Proteins will appear as clear zones in a translucent blue background. The gel can be completely de-stained by repeated washed in the de-stain solution.
N.B. Equilibrate the gel in transfer buffer before proceeding with the blotting.
Transfer can be done in wet or semi-dry conditions depending on the blotting apparatus.
However, the principle is the same in each case. Just as the proteins with an electrical charge (provided by SDS bound to them) can be induced to travel through a gel in an electrical field, in the same fashion the proteins can be transferred in an electrical field from the gel onto a membrane that “blots” the proteins from the gel.
Semi-dry transfer is faster, however wet transfer is generally less prone to failure due to the drying of the membrane and is especially recommended for larger protein >100KDa.
N.B Remember to always wear gloves when dealing with proteins because hands are rich of proteins.
To perform a Western blot starting from the end of the electrophoresis, remove the glass plate and transfer the gel into Transfer buffer. Equilibrate the gel between 10-30min at room temperature on a shaker according to specific protocols. Equilibration facilitate the removal of electrophoresis buffer salts and detergents. Salts, if not removed may increase the conductivity of the Transfer buffer resulting in increased heat during the Transfer. Also equilibration allow the gel to adjust to its final size (acrylamide gel will shrink in methanol-containing buffer) thus improving the efficiency of the transfer,
At the same time cut the membrane of the same size of the gel and equilibrate in Transfer buffer as well. The two most used types of membrane for proteins are nitrocellulose and PVDF membrane. If you are using PVDF membrane remember that is essential that you pre-wet the membrane in 100% methanol before immersing it in Transfer buffer.
The above steps are necessary for both wet or semi-dry transfer.
The main difference is the assembly of the Blotting system.
Both the pads and the Blotting paper should be pre-wet in Transfer buffer.
Once assembled, insert the cassette in the transfer cell (Fig1. B) making sure the black side of the cassette is facing the black side of the transfer cell. In this case the “sandwich” is submerged in Transfer buffer to which an electrical field is applied (i.e. 250mA for 1hr. Always check specific protocols.). Since the current will produce high heat always pre-cooled the Transfer buffer and use the cooling pack provided with the kit.
Alternatively, the semi-dry Western blot apparatus (Fig.2) should be assemble as follow: starting from the platinum plate (anode) / Blotting paper / membrane / gel / Blotting paper / stainless steel cathode lid / safety cover
Usually mini-gels are transferred for 15-30 minutes at 10-15V. Never exceed 25V even for large gels.
Visualisation of protein on membrane
N.B. This step can be omitted
It is possible to visualise the proteins on the membrane to check for successful transfer using Ponceau Red.
Wash the membrane briefly in 1xTBS-Tween20 (TBS-T); alternatively, PBS based buffer can also be used throughout the all procedure. Incubate the membrane in Ponceau Red for 5 minutes and then rinse extensively in water until the protein bands are well-defined.
The membrane can be completely de-stained by washing several times with 1x TBS-T. If using PVDF membrane re-activate in methanol and then wash in 1xTBS-T.
Blocking the membrane
Blocking the membrane prevents non-specific background binding of the primary and secondary antibodies to the membrane.
The most commonly used blocking buffer are non-fat milk and BSA. The milk is cheaper but is not recommended for the study of phospho-proteins (milk contains casein which is a phospho-protein)
Blocking time can vary between 2-3 hours at room temperature on a shaker to 24hrs (overnight blocking should always be done at 4oC)
Incubation with primary antibody
After blocking the membrane should be washed 3x for 10 minutes in TBS-T
Primary antibody should be diluted in fresh Blocking buffer at the recommended dilution and the blot incubated at room temperature for 2 hours on a shaker (it is possible to incubate overnight at 4oC).
Incubation with secondary antibody
Prior addition with secondary antibody the membrane should be washed 3x 10 minutes in TBS-T
Secondary antibody should be diluted in Blocking buffer at the recommended dilution. Incubation time vary between 1 to 2 hours at room temperature shaking (never incubate secondary antibody overnight).
Wash the membrane 3x 10 minutes in TBS-T and 1x in TBS (no addition of Tween20)
There are several choices of readout for Western Blotting according to the molecule conjugated to the specific antibodies:
Western blotting with colorimetric detection uses secondary antibodies conjugated to an enzyme which catalyses the conversion of a soluble chromogenic substrate to a coloured insoluble product that precipitate onto the membrane to produce coloured bands that can be visually observed. For example, for ALP-conjugated antibodies BCIP/NBT detection kit is generally used.
Developing the blot is simple: the membrane is incubated with the chromogenic substrate until the required level of signal is developed, then the substrate is simply washed away. This stops the enzymatic reaction and halts the blot from developing further.
Colorimetric detection is quick and easy and doesn’t required a specialised system for imaging, however it is not possible to strip and re-probe the membrane if required. Moreover, it is possible to increase the sensitivity of the signal but prolonging the incubation, but this can lead to increased background noise which can obscure the signal for the protein of interest. Therefore, this method is best suited for abundant proteins.
Fluorometric detection requires the use of antibodies which have been labelled with a fluorophore. A light source is used to excite the fluorophore, which then produces a transient light emission as it returns to its ground state. The light is emitted at a higher wavelength than that which was used for excitation and is detected with specialised readers such as the chemiPRO Imaging systems series. Advantages to use this detection method includes:
- Multiplexing Use of multiple fluorophores for simultaneous detection of several target proteins making stripping and reprobing unnecessary
- Accuracy Fluorescent detection offers a more accurate quantitative method than an enzyme-based one which relies on a chemical reaction
- Stability Most fluorescent molecules provide excellent stability, allowing blots to be archived and re-imaged later on.
Chemiluminescent Western blotting is a highly sensitive protein detection method. The broad dynamic range allows analyte detection over a wide range of protein concentration. Chemiluminescence detection occurs when a substrate is catalysed by an enzyme and produce light as by-product of the chemical reaction. For example, for the detection of HPR-conjugated antibodies, ECL are the substrates generally used. The light is then captured using a cooled CCD camera such as the one found in our chemiPRO and chemiLITE Imaging systems.
The linear response of reporter enzymes combined with enhanced chemiluminescent substrates makes the detection method suitable for quantitative assays. The signal will decay upon exhaustion of the substrate, though the enzyme will remain active. The signal can be refreshed by adding new substrate if required.
Proteins perform many critical functions in everyday existence for any cell and organism. Therefore, to better understand how, when and where a specific protein performs its specific functions (i.e. enzymes, structural proteins, cell membrane receptor, etc.) or what might affect that protein to change its behaviour, it is fundamental to be able to identify that one protein in a complex mixture of other proteins.