We use western blotting to detect specific proteins from a sample and to gain information about the size and the relative and absolute amounts of these proteins (see Quantitative Western Blotting). The technique involves the electrophoretic separation of denatured proteins followed by blotting and transferring them to PVDF or nitrocellulose membranes. Next, the proteins are renatured and the blots are incubated with primary antibody followed by horseradish peroxidase conjugated secondary antibody. Finally, they are detected using the electrochemiluminescence method.
Preparing Samples for Western Blotting
In a 1.5 mL microcentrifuge tube prepare 1 mL of 1X loading buffer
by combining:
250 µL 1M DTT
500 µL 2X loading buffer
250 µL distilled water
Obtain samples and place in 1.5 mL microcentrifuge tubes.
Dilute samples by adding at least an equal volume of 1X loading buffer.
You can add enough 1X loading buffer to make a maximum of 25 µL in total.
Preparing Samples for Quantitative Western Blotting
Prepare 1X loading buffer, and dilute samples with it as described above.
Also prepare serial dilutions of a known quantity of the protein of interest. This will allow you to determine the absolute amount of protein in your samples.
You should have your tank already prepared with the electrophoresis module and cathode and anode buffers.
Fill the tank
with running buffer to just under the edge of the outer gel plate.
Load samples into the wells with a Hamilton syringe.
Cover the tank and connect the electrodes.
If not already done, program running conditions
Run gels for 40 min @ 30V (constant voltage) to stack proteins then 1 hr @ 40-50mA (constant current) to separate proteins
After electrophoresis is complete, turn off the power supply and disconnect the leads.
Remove the tank lid, lift out the assemblies and discard the running buffer.
Remove the gel by inverting the gel and plate under fixative or transfer solution and agitating it gently until it separates from the plate.
Rinse the module with distilled water after use.
Electrophoretic Separation for Quantitative Western Blotting
Perform electrophoretic separation as described above.
Load the serial dilutions prepared above on the same gel as your samples.
Transfer
(refer to Trans-Blot ® SD Semi-Dry Electrophorectic Transfer Cell Instruction Manual)
Remove gels from apparatus and presoak them in transfer buffer for 15 min.
Obtain nitrocellulose membrane. If proteins are hydrophobic, use PVDF membrane instead.
Cut the membrane to the dimensions of the gel.
Soak the membrane in transfer buffer for 15-30 minutes.
Cut filter paper to the dimension of the gel. One piece of extra thick filter paper or 3 pieces of VWR Blotting paper #703 per gel are needed -for each gel/membrane sandwich. Completely saturate the filter paper by soaking in transfer buffer.
Assemble the Semi-dry electrophoretic transfer cell by removing the safety cover and the stainless steel cathode assembly.
Place a pre-soaked sheet of extra thick filter paper onto the platinum anode. Roll a pipet or test tube over the surface of the filter paper to exclude all air bubbles. Repeat with one or two more sheets of buffer-soaked filter paper.
Place the blotting paper on top of the filter paper. Roll out air bubbles.
Place the gel on top of the transfer membrane. Roll out air bubbles.
Place one thick sheet or three thin sheets of soaked filter paper on top of the gel.
Place the cathode onto the stack. Press to engage the latches with the guide posts.
Place the safety cover on the unit & plug into power supply red wire to red outlet and black wire to black outlet.
Turn on power supply and transfer at 50mA per gel for 1 hour.
Following transfer turn off power supply and disconnect the unit from the power supply. Remove safety cover and the cathode assembly. Discard the filter paper.
Stain the gel to determine transfer efficiency, or use pre-stained molecular weight standards. Or, stain a portion of the membrane with colloidal gold, Biotin Blot Total Protein Stain, or an anionic dye such as Amido Black. Zeta-Probe membrane can be stained with Biotin-Blot Total Protein Stain.
Place nitrocellulose into approximately 20 mL of 1X Blocking buffer
Wash and incubate
Block membranes in 1X blocking buffer at room temperature for 30 minutes.
Incubate blots overnight at 4°C in primary antibody diluted in either polyclonal antibody buffer or monoclonal antibody buffer , depending on the antibody utilized.
Wash the blots once in 1X wash buffer followed by two additional washes in 10X TBST.
Incubate the blots in horseradish peroxidase conjugated secondary antibodies diluted in TBST with 1% BSA for 2 hours at room temperature.
There are a number of important considerations if you want to use western blotting quantitatively. The first is to remember that a lot of what people tell you is ill informed. There are camps out there that say you cannot do western blotting quantitatively - just because they can't do it does not mean you cannot. More often people just blindly quantify any old rubbish and believe the densitometer even when it the data doesn't make sense. Both usually means they have not thought through carefully what it is that they are doing.
ECL is an enzymatic reaction that gives off light. That means for the light coming out to be linearly proportional to the amount of enzyme present you need a first order or psuedo first order reaction. To get that you need to be in substrate excess. This is not always the case when developing western blots. Usually one has a limited amount of reagent added as a film across the top of the blot separated from the photographic film or the camera but a thin plastic layer. This means that it is quite easy to use up a lot (or all) of the substrate on your band without using up substrate on faint bands. As a result the response will not be linear and your quantification will be wrong. The extreme example of this is when some bands are 'white', 'clear' in the center but black around the edge. This is because there is no subtrate left in the center of the band. There are lots of other areas where non-linearity creeps into your experiment. The next most common (and most difficult to control) is the way your protein binds to the membrane. The critical issues are how well the protein binds and the binding capacity of the blotting material. Another source of error is the linear range of the film - although this is much less prone to error than most people think. The response of good film to light is linear over three orders of magnitude. Usually protein binding issues limit the membrane to 1-1.5 orders of magnitude. So you are almost always limited by the membrane and your protein. However - it is important to remember that black is black and you cannot get blacker than black so a band intensity 10X darker than a black band is still black - ie it reads the same.
Solutions: First - it is a good idea to put the substrate on right before you develop the blot. Make sure there is enough reagent. Sometimes putting a piece of filter paper under the blot and soaking it with substrate helps. Second - just like in other protein assays a standard curve is critical. Both were used in the example below. In this example I have used one of the more difficult quantifications. It 'enjoys' two other problems. A GST-fusion protein is being used as standard - to do this you must be sure that there are the same epitopes in the GST-fusion protein as in the protein you intend to measure. There are some cross reacting bands. We have assumed that they are not degraded GST-fusion protein.
Calulate the absolute amount of protein in the sample:
Confirm purity of the standard. In Figure A an SDSPAGE gel of 1 ug of purified GSTBcl2ΔTM was determined to be >95% pure by qunatifying the Coomassie blue stained gel shown in A using a densitometer
Use the serial dilutions (figure B) of the standard to create a standard curve on the same SDS-PAGE gel as sample lysates expressing the protein of interest. Standard curves must be on the same blot to account for transfer problems and the amount of reagents, development time etc.
Obtain “net intensity” readings of the serial dilutions of the standard using Kodak Image Station. Alternatively use film and a standard scanner. In practice there is little difference between the techniques. Use these to create a standard curve.
Perform a linear regression using MS Excel software (Figure C). Only use lines of best fit with R 2 values ≥ 0.95, to ensure that the readings are within the linear range of the equipment. - Visually inspect the line of best fit - Never just trust the calculations. You would not believe the number of times I have found missing decimal points or calculations pointing to the wrong cell in an excel spread sheet.
Compare "net intensity” readings of the sample lysates to the standard curve to calculate the amount of protein in the unknown samples (see Figure D).
Notice how tight the error bars are when you do it right! (the data comes from 3-5 completely independent experiments and is Std. Dev.
