Western blot (WB), also called Immunoblot, uses specific antibodies to analyze one protein in a sample containing multiple proteins. Proteins are separated from each other by size using gel electrophoresis, then transferred to a membrane and detected with a specific antibody. This technique is widely used in biological research, as it provides information about the molecular weight of a protein as well as relative differences in expression between samples.
Review advice on sample preparation and troubleshooting tips in this webinar.
Caitlin Buckspan received her B.S. in Biological Engineering from the University of Florida in 2007 and a M.S. in Bioengineering from the University of California, San Diego in 2010. At the University of Florida, she researched seasonal changes in termite hormone levels, where her research relied heavily on western blots.
Caitlin’s Master's thesis focused on age-related changes in mesenchymal stem cells using a murine model. She joined the Abcam Scientific Support Team in 2010, and is a current member of the Histochemical Society and the International Society for Stem Cell Research.
JR: Hello, and welcome to Abcam's webinar on an Introduction to Western Blot Principles and Troubleshooting. The principal presenter will be Caitlin Buckspan, a member of Abcam's Scientific Support Team. Caitlin received her B.S. in Biological Engineering from the University of Florida in 2007 and a M.S. in Bioengineering from the University of California, San Diego in 2010. At the University of Florida, she researched seasonal changes in termite hormone levels, and this research relied heavily on western blots. Her Master's thesis focused on age-related changes in mesenchymal stem cells using a murine model. She joined the Abcam Scientific Support Team in 2010, and is a current member of the Histochemical Society and the International Society for Stem Cell Research.
Joining Caitlin today will be Mandeep Sehmi, a member of our Core Focus area team. Mandeep completed her Bachelor's Degree in Applied and Human Biology at Aston University, and an MSc in Molecular Pathology and Toxicology at the University of Leicester. Mandeep went on to work at the Medical Research Council Toxicology Unit in Leicester, working on B-cell leukemia and completed a PhD in B-cell Immunology in cancer during her time there. In her PhD, she worked on identifying a novel membrane protein in B-cell lymphoma called HVCN1, which was shown to have an important role in normal B-cell function, and then subtypes of B-cell lymphomas. Mandeep joined Abcam in 2011.
If you have any questions during the presentation, they can be submitted at any time through the Q&A panel on the right hand side of your screen. Your questions you submit will be answered during the troubleshooting section at the end of the webinar. I will now hand it over to Caitlin, who will begin her presentation on western blot principles and troubleshooting. Enjoy the webinar!
CB: Hi everyone, thanks for joining us for our western blot webinar. Today we'll be going through the basic principles of the western blot, and I'll show some examples of troubleshooting this technique. Remember that if you have any questions, please submit them to the Q&A panel on the right hand side of your screen, and I'll be available to answer some of them at the end of our presentation.
So what is a western blot and how does it work? It's a complicated technique, but to summarize it's used to identify a specific protein of interest within a sample containing many proteins. First, the proteins are separated from each other according to molecular weight using gel electrophoresis, and then a specific antibody is used to identify the protein of interest. So we can determine the molecular weight of the protein of interest by comparing the results to protein standards of millimolecular weight, and we can also determine the relative quantity of the protein of interest among samples. For example, in this western blot image, you can see that the band of interest, which is the control protein beta actin, is around 38 kDa we compared to these proteins of known molecular weight, which are run side-by-side with our samples. In lane two, less antibody was used to stain this particular sample, and so the band appears fainter than the others.
Basically, western blots are useful tools any time you'd like information about protein expression in your samples, such as those listed here. They can be used to indicate whether the sample expresses the protein of interest at all, or they can be used to give information about relative expression among different samples or treatment groups. They're awfully useful to validate an antibody specificity if your end goal is to use it in immunohistochemistry, or immunocytochemistry as cross-reaction can be easily seen in western blot, since proteins are separated from each other visually.
In this presentation, we'll go over the steps of the western blot and talk about controls that should be used during your experiment, as well as the selection of an appropriate antibody for your assay. I'll end with several troubleshooting and optimization examples of common problems in western blotting. The first step to a great western blot is preparing your sample correctly, because the final result will never be of better quality than the starting material. There are three main steps to preparing your sample: lysis, measurement of protein concentration and reduction and denaturation.
Lysis, essentially solubilizes, the proteins from the cells or tissue using buffers and mechanical agitation. All of this is done on ice with the addition of fresh protease inhibitors to the buffer, because keep in mind that there are active protease enzymes in cells and tissues that can break down your sample, if allowed. The protocol is slightly different for cells or tissue samples, but it's really the same concept. Homogenize the sample in a lysis buffer and agitate, then centrifuge and collect the supernatant.
Once the samples are fully lysed, measure the protein concentration. You can use a Bradford assay or BCA assay, whichever method you prefer. This will be important to know for the electrophoresis step, in order to make sure you are loading equal amounts of protein per sample.
The final step of sample prep is reduction and denaturation, this breaks down higher level structures so that proteins can be separated during electrophoresis, according to their primary amino acid structure. This is done by adding a buffer containing a reducing agent, as well as detergent and heating generally for 5 min around 95°C. Heating at a lower temperature may be more preferable for some proteins, particularly multi-pass membrane proteins, which can aggregate at higher temperatures.
Next, we'll discuss lysis reduction and denaturation a little further. So a few more suggestions for the lysis step, and this is important to make sure proteins are actually fully solubilized. First, as we discussed, keep everything cold and on ice as possible, and add protease inhibitors fresh to the lysis buffer. Additionally, if you plan to detect a phosphorylated form of a protein, you'll also need to add ample phosphatase inhibitors. Finally, the optimal lysis buffer depends on the cellular localization of the protein of interest, as a stronger buffer like NP-40 or RIPA is needed to fully solubilize membrane or organelle-bound proteins from the cells.
For reducing and denaturing in sample buffer, the buffer contains two agents: SDS or sodium dodecyl sulphate, and either beta-mercaptoethanol or dithiothreitol, abbreviated DTT. The SDS is a denaturing agent which helps break the protein down to its amino acid structure, and it also coats the protein in negative charges at a uniform mass ratio. This uniform ratio of negative charges will be essential during the next step of the procedure, gel electrophoresis. Beta-mercaptoethanol and DTT are reducing agents which break their sulphide bonds, also breaking down the protein to a linear primary amino acid structure, which is all simplified in this image. As a side note, occasionally the protein of interest will need to be non-reduced or non-denatured, often because a particular antibody only recognizes this form of the protein. So the reducing and denaturing steps can be skipped, if needed, but most western blots are run and denatured reduced conditions. So once the samples are properly prepared they're ready for gel electrophoresis.
We've mentioned that gel electrophoresis is used to separate proteins according to molecular weight. We're actually separating the proteins in an electric field and taking advantage of the fact that different sized proteins will move through a gel matrix at different rates. This image shows a typical apparatus used for gel electrophoresis.
The type of electrophoresis most commonly used in western blotting is abbreviated SDS-PAGE. SDS is the denaturing agent which is added to the buffer during this procedure. PAGE stands for polyacrylamide gel electrophoresis, as the gels are typically made of polymerized acrylamide. So how does SDS-PAGE work? We start with a porous gel matrix made of highly cross-linked acrylamide, submerged in a chamber filled with an aqueous buffer. Samples of protein lysates are loaded into wells of the gel. The well is the top of each lane, which is basically a depression in the gel, so there's space to hold the lysate until the current is applied. When the current is applied, the proteins will migrate towards the positive pole, as they are coded in negatively charged SDS. Because smaller proteins are less inhibited by the gel matrix, they will move faster through the gel, and when the current is removed they will be farther down the gel, so proteins are separated from each other according to size.
Gels are made of polymerized acrylamide and cross-linked by Bis in the presence of ammonium persulfate and TEMED. The structure of the gel can be controlled by altering the acrylamide percentage. Basically, by increasing the total acrylamide percentage, the pores of the gel matrix become tighter, which slows down small proteins. For this reason, high percentage gels are great when studying smaller proteins, and vice versa, a lower acrylamide percentage results in larger pores of the gel matrix, which allows larger proteins to migrate more easily and separate more efficiently in the gel. There are also gradient gels consisting of layers of different acrylamide percentages, so that those small and large proteins can be efficiently run on the same gels. I would recommend these if you're analyzing a large range of proteins on the same blot, or if you're unsure of the molecular weight of your protein of interest.
As mentioned previously, sometimes you don't want the protein of interest to be in the reduced denatured form. This will affect the recipe of your running buffer during electrophoresis, as well as your loading buffer during sample preparation. Here is a chart that can be used as a reference, but basically you can remember that if the protein needs to be in the non-denatured, also called native form, remove SDS from the loading buffer and the running buffer. If the protein needs to be in the non-reduced, also called the oxidized form, remove the beta-mercaptoethanol or DTT from the loading buffer.
The next step after gel electrophoresis is protein transfer. The transfer step is where the word 'blot' in western blot originates, because the proteins are going to be transferred out of the gel onto another surface. This is because gels are a poor substrate for immunodetection. Antibodies don't stick to the proteins and the gels are very fragile, and can be easily torn. So we transfer the protein to a more durable substrate for subsequent immunostaining.
How does Transfer work? It's a similar principle to electrophoresis, and that will move the negatively charged proteins towards a positive pole in an electric field. The gel is placed adjacent to a porous membrane, and this sandwich of membrane and gel is added to a transfer cassette with the membrane closest to the positive pole. Be careful to avoid air bubbles between the gel and membrane, as air is an insulator and won't conduct a current, so proteins will not transfer through the bubble. I suggest rolling the transfer sandwich with the long edge of a conical tube to remove air bubbles. Once the sandwich is in the cassette, an electric field is applied and the proteins migrate and adhere to the membrane in the same configuration as they are in the gel.
There are many different kinds of membranes available, there are different kinds of material, commonly PVDF or nitrocellulose, and there are also different pore sizes. Each has different characteristics, I suggest using PVDF when blotting for proteins that are difficult to transfer, but PVDF can also contribute to high background in some cases. So you can try a few different membranes and see what works best with your system, and with your sample protein. After transfer, I always recommend using a reversible protein stain such as Ponceau red on the membrane to ensure that the proteins fully transfer from the gel.
There are two common methods of transfer: wet and semi-dry. The main difference being that in wet transfer the sandwich is submerged in the chamber filled with transfer buffer, and in semi-dry the sandwich is placed directly between the cathode and the anode. Semi-dry is faster, but the membrane can dry out and inhibit protein transfer. Wet transfer is more time-consuming, but it's the optimal method to use when transferring a protein that might be difficult to transfer, such as a large protein or a hydrophobic protein.
Speaking of transferring large proteins, there are some steps I recommend when transferring proteins that are larger than 100 kDa. Use wet transfer and transfer overnight at low voltage and low temperature. Also, add SDS to the transfer buffer, which will help prevent the protein from precipitating in the gel. Reduce methanol to 10% or less in the transfer buffer, which helps the gel swell and release proteins. If you're using a PVDF membrane, methanol can be removed entirely from the buffer. When transferring large proteins it will also be important again to use a Ponceau stain on the membrane to make sure that the protein did indeed transfer. After the protein has transferred to the membrane, it's ready to be stained and visualized with a specific antibody.
Why do we need antibodies? You could use a protein stain like Coomassie in a gel, or Ponceau on the membrane, but these are non-specific stains and will show every protein in the sample, such as the image on the left. It's very difficult to tell which protein is the protein of interest, and it would be even harder to attempt to compare the quantity of that protein between samples. So, specific antibodies are used so that only the protein of interest is visualized, as in the image on the right.
The first step in the immunostaining procedure is to block the bare space on the membrane with a neutral protein. This is important because the membrane is an adherent surface for all proteins, so a free antibody could non-specifically bind to the exposed membrane, but antibodies shouldn't bind to a neutral protein. To block the membrane, the membrane will be placed in a dish and covered with a solution of TBST and 3-5% milk or BSA. The proteins in the milk or the BSA bind to the membrane to cover the free membrane, so that antibody won't stick to these areas later. The dish should be placed on a rocker to agitate for 1 hr at room temperature, or overnight at 4°C.
Once the membrane is thoroughly blocked, pour off the blocking solution and add the diluted primary antibody solution to cover the membrane. The antibody can be diluted in TBST or in the blocking solution. If you're using the antibody for the first time with your sample, you may want to try a few different dilutions of antibody in order to find the optimal dilution factor. Leave this to incubate for an hour at room temperature or overnight at 4°C on a rocker, and the antibody will bind to the protein of interest on the membrane.
Once the antibody has bound, the membrane needs to be washed to remove any residual antibody. I would recommend 3-5 washes with TBST on a rocker for around 10 min each wash.
Following the primary antibody, the membrane will be treated with the secondary antibody that is conjugated to a molecule that can be easily visualized. The secondary antibody will be specific for the primary antibody, so that it binds to the primary antibody, and thus is extensively bound to the protein of interest. Leave the secondary antibody to incubate for around 1 hr at room temperature.
Again, wash the membrane thoroughly to remove residual secondary antibody. 3-5 times with TBST for around 10 min should be sufficient. Once the membrane is washed it is ready to detect the protein via the molecule conjugated to the secondary antibody.
There are three kinds of detection that are most commonly used with the western blot. The first is detection using a chemiluminescent substrate, such as ECL or ECL Plus, which is more sensitive. The secondary antibody in this case would be conjugated to HRP, horseradish peroxidase, which catalyzes a reaction with the ECL to produce a luminescent signal which is captured on film. Similarly, a chromogenic substrate such as 4-chloronaphthol or DAB can be used with an enzyme conjugated secondary antibody, to deposit a colored product directly on the membrane. In this case, there is no special detection equipment needed, so it's a good choice if you're just starting to run western blots in your lab. A third option is to use a secondary antibody conjugated to a fluorescent molecule, and this fluorescence can be detected using a specialized detection system.
As part of every well-planned experiment, there are certain controls that should be used and now we'll talk about some of the important controls to use in western blotting. A loading control is often used during western blotting and it's an antibody that is used in addition to the antibody directed against the protein of interest. The loading control targets a common protein that samples should contain in roughly the same proportion. So that assuming the same amount of protein from each sample was loaded correctly to the gel during electrophoresis, there should be similarly sized bands of the loading control target. If there was a mistake during loading, the loading control band will show this error. The target that you choose will depend on what kind of sample lysate you're using. If you have a whole cell lysate, generally beta actin, GAPDH or tubulin will be used as a loading control. For a mitochondrial fraction use VDCA1 or COXIV, and for a nuclear fraction use a lamin B1 or a TATA binding protein antibody.
One important note of caution is to be careful when comparing loading control results between dissimilar samples, as an antibody's binding affinity can vary among different forms of the protein. In this image, for example, each lane is loaded with the same quantity of the sample from a different species, and stained with a beta actin antibody. Although the lanes are loaded correctly, the band in the fish samples in lane three is fainter than the others, but this is because the antibody has a lower binding affinity for the fish beta actin protein.
In addition to a loading control, it's a good idea to run a positive control and a negative control sample alongside your unknown sample. The positive control is basically a sample known to contain the protein of interest, such as a recombinant protein, a transfected cell line, or tissues or cells known to express the protein. A negative control is known to not contain the protein of interest, which could be a knockout or siRNA sample, or tissues or cells that don't express the protein of interest.
Another useful control is the blocking peptide, which is more commonly used with antibodies in which the immunogen is a short peptide sequence, as opposed to a full-length protein. To use the peptide as a control, the antibody is pre-treated with the peptide so that the specific binding sites of the antibody are bound and blocked by the peptide. Then the pre-treated antibody is incubated with the membrane. Any specific bands will not appear on the membrane, since the binding sites are blocked. If any strong bands do appear, they must be due to non-specific binding of the antibody. In this image there are no bands in lanes three and four, and this is because the antibody used on these lanes was pre-treated with the immunogenic peptide. In the other lanes the antibody was pre-treated with peptides very similar to the immunogen, and since bands appear in these lanes, it demonstrates that the antibody is not binding to the non-immunogen peptides, showing the antibody's specificity.
Next, we'll go over a couple of things to keep in mind when selecting an antibody appropriate for your western blot. The first criteria, which might seem obvious, is to choose an antibody that has been tested successfully in western blot. This is important, because not all antibodies will work well in a western blot. If you're on the Abcam website, we will list WB in the tested application section if we have tested the antibody and it works in western blot. So just make sure to check that section when looking for a suitable antibody. If western blot is not listed as an application, the antibody was probably not tested yet in this application, and we encourage you to contact our scientific support department in this case. The contact details for our scientific support team will be given later in this presentation.
Now, the specific region of a protein that an antibody targets may or may not be important to your study. If you're studying a specific isoform of the protein, or the precursor versus the mature protein, you should be interested in the specificity of the antibody. You can often find out which region the antibody targets by looking at the immunogen sequence that the antibody was raised against, as the animal will produce antibodies that recognize this particular sequence. If the antibody is polyclonal, then it will recognize the entire immunogen sequence, but if it is monoclonal it will only recognize one short part of the sequence. If there is a specific isoform of a protein that you are studying, make sure the immunogen region overlaps with the region that your specific target contains.
So now that we've covered how to run a great western blots, we'll talk about how to troubleshoot the results when they aren't as great as expected. You might eventually run a western blot that looks something like this image where the lanes are completely blank, or there might be faint bands of the expected molecular weight. If there are no bands at all on the blot, the protein might not have transferred to the membrane, which can be checked again with a Ponceau stain, or maybe the secondary antibody is not compatible with the primary antibody. It's possible that the sample does not express the protein of interest, which can be checked using a positive control sample side-by-side with your unknown sample. If the bands are very weak, you can increase the amount of protein loaded and the concentration of the antibodies used. You can change the blocking agent or block for a shorter period of time, and you can increase the times of the antibody incubations. If you're using a chemiluminescent detection, I would also suggest exposing the film for a longer period of time to pick up any bands that might be on the blot.
You could also run into an issue of high background or more bands on the blot than expected. I would recommend increasing the amount of lysate loaded onto the gel, diluting the antibody out further, changing the blocking agent and try a nitrocellulose membrane if you're using PVDF. You can also decrease the exposure time if you're using a chemiluminescent detection. If there are extra bands at lower molecular weights than the protein of interest, degradation may be to blame. Make sure protease inhibitors are added fresh to the sample buffer, and that the samples are kept on ice. If there are extra bands of molecular weight that are multiples of the expected molecular weight, such as at 100 kDa when there should only be bands at 50 kDa, boil samples a bit longer to make sure any multimers are disrupted. You can check the literature, and I recommend the SwissProt website to see if the protein is modified post-translation, such as with glycosylation, acetylations or phosphorylations, or if it has different isoforms or cleavage fragments, which could explain the extra bands on the blot.
Here is a specific example from our own in-house testing, which there is very high background on a blot with a new lot of an antibody. The technician thought they may have incorrectly diluted the antibody which was intended to be at 1 mg/ml, so they simply remade the working dilution from the stock solution, and they also diluted the sample lysate 10 times and 100 times just to be on the safe side. The new results showed an ideal western blot, just from diluting the antibody and sample.
These are two other interesting pieces, as these antibodies work much better with a specific blocking agent. The antibody on the left, as you can see, shows the multiple bands when the membrane is blocked with 5% BSA. However, a clear, yet faint, band is seen using 5% milk as a blocking agent. This isn't to say that milk is always the ideal choice though, the antibody in the right image gives a much stronger signal using 5% BSA and it's barely detectable when the membrane is blocked with milk. Many antibodies will work just fine with either milk or BSA, so you can experiment with different blocking agents if the initial results are not optimal. If you're blotting for a protein using a phospho-specific antibody, meaning that it should only detect the protein if it's phosphorylated. We suggest using BSA as a blocking agent. Milk endogenously contains phosphatases which can cleave the phosphate group from the protein of interest, so that the antibody cannot bind, resulting in no bands. If we do know that a particular antibody works better with one blocking agent, we will put that directly on the datasheet, but always feel free to contact us about an antibody if you'd like more details about how it's been tested.
Here is an example of a blot with multiple problems, and I'll just point out that this antibody did not pass QC and is not for sale, so don't worry. But this image does show some common problems that I'd like to discuss. First, there is evidence of air bubbles that were present during the transfer step, which did not conduct the current during transfer, and so protein did not migrate to the membrane at this spot. An easy way to avoid this is to roll out the gel and membrane sandwich using the side of a conical tube while it's being assembled. There are also some spots of poor washing where a reagent has pooled on the membrane. To avoid this make sure all solutions cover the entire membrane, and rock or agitate the membrane during incubations. The last thing I'd like to point out is the area in the right upper corner where there was incomplete contact between the gel and the membrane, and this may have been due to folding of the gel or tearing of the gel. If the gel isn't flat on the membrane, again, smooth out the sandwich with the side of a conical tube to make sure all areas are in contact. If your gels frequently tear, you can try using a different kind of gel such as the Optiblot gels that we've recently added to our catalogue, and these are up to 10 times stronger than traditional gels. So there are lots of things that can go wrong during a western blot, but these are fixable problems. It's a good habit to stain your membrane with a Ponceau red stain after transfer, as you can catch these problems early and rerun the electrophoresis and transfer, if necessary.
Here is another example of incomplete transfer, but in this case only the large proteins are having trouble being transferred to the membrane. You can see that there should be at least a high molecular weight standard of proteins, but these have not efficiently transferred. If you see something like this, simply follow the suggestions mentioned before for transferring large proteins, such as a wet transfer overnight at low voltage and lowering the amount of methanol in your transfer buffer. If you do see the reverse problem, meaning only high molecular weight proteins are on the membrane, you may have over-transferred so that the small proteins migrated straight through your membrane, so shorten your transfer time or voltage in that case.
Finally, for the last example, here's the case of an antibody that has much higher affinity for the protein in its non-reduced, non-denatured conditions. Meaning that SDS and beta-mercaptoethanol or DTT, are not used in a loading buffer or PAGE-running gel. This image was submitted as part of an excellent AbReview by an outside researcher, who found that a collagen III antibody gave a weak band when the protein was reduced and denatured, seen in the left lane. Stronger bands were seen when the protein was not reduced and denatured in the right lane sample. The band is around 300 kDa, is a dimer made of two alpha chains that individually would run around 130 kDa in reduced, denatured conditions. In its native form the protein has a three-dimensional shape and the antibody likely has a strong bonding site along the 3D shape, but is lost once the protein is reduced and denatured.
On a final note, I'd like to reiterate how important it is to really understand the biochemistry of your protein of interest, as western blotting depends heavily on this knowledge. Each blot would not necessarily give you one band at the expected molecular weight, but this isn't necessarily a cause to worry. There may have been post-translational modifications or cleavage events, which cause the protein to run at a different size, or your sample may express a particular isoform of the protein. When unsure, consult resources such as SwissProt website or the available literature for more information about your target, and always feel free to contact the Abcam scientific support team if you need any assistance with this or with any part of your western blot. Contact details for the scientific support team will be provided later in the presentation.
Now we'll turn the presentation over to Mandeep, who will give an overview of the research tools that Abcam provides for your western blotting needs. All of our sites will be available to download at the conclusion of the webinar for your reference. Following Mandeep's presentation, I'll be available to answer some of the questions that have been sent in to us, so please continue to submit questions, if you haven't already.
MS: Thank you, Caitlin. As I mentioned, I am going to review some of the products Abcam offers to support your western blot experiments. Before I do so I would like to remind you to feel free to submit any questions you would like Caitlin to answer. In addition to well-characterized, primary and secondary antibodies, our catalogue also includes a number of products to support your western blotting from our reagents range to our MitoSciences range. To begin with, I am going to introduce our reagents range, which includes our core western blotting range, Optiblot, our prism protein ladders, EasyLink conjugation kits, as well as controls and lysates, and ECL kits. As Caitlin mentioned, a PDF of the slides will be available for download after the webinar, so you can refer back to these links.
Our Optiblot range includes gels, buffers and accessory reagents which are designed to enhance your western blotting and improve on existing products on the market. Using pre-cast gels from our Optiblot range, you can benefit from a one-year shelf life, as well as other features that make the gels easier to use, such as wells that are easy to visualize and increase strength and ease of use. To accompany the pre-cast gels, we also offer buffers that are formulated for optimal running of the gels, which can be purchased or prepared in the lab using buffer recipes available on our website. Also, our Optiblot range includes accessory reagents such as Optiblot Blue, allowing you to stain proteins in 15 min; an Optiblot Bradford reagent, for detergent compatible protein quantitation. We also offer reagent bundles which include all gels and buffers you will need to conveniently run your experiments. You can find additional information about the Optiblot range by visiting the link shown.
Our reagents range also includes pre-stained prism protein ladders, which can be used for estimating the size of your protein, and determining western blot transfer efficiency. With multi-colored fluorescence bands you can easily estimate the size of your protein, and their broad range covers a wide range of molecular weights. If you would like to carry out a more rapid detection using a primary antibody, you can use our EasyLink conjugation kits to conjugate a label such as HRP or alkaline phosphatase to your primary antibody in only 20 min. Additional to western blotting accessory reagents, also include loading controls to ensure equal amounts of lysate and loading onto your gel. Lysates which can be used as positive controls and ECL kits for sensitive detection. Please visit the links shown to explore these product ranges further.
If you are working in metabolism or mitochondrial research, Abcam recommends our MitoSciences range of products, which offer a comprehensive selection of monoclonal antibodies, lysates and kits designed to optimize your research. MitoSciences monoclonal antibodies are highly optimized and validated to work in a range of species. These include human, drosophila, monkey, cow, rodent and yeast. We also have mitochondrial lysates isolated from specific tissues which can be used in your western blotting as positive controls. If you would prefer to isolate a mitochondria or cytoplasmic proteins yourself, we also offer a range of tools for cell fractionation.
Our popular western blotting antibody cocktails are optimized, pre-mixed solutions of antibodies for western blot analysis of key panels of targets and control enzymes or proteins. These cocktails offer the advantage that a number of proteins can be detected in one lane, so you can analyze a variety of samples in one western blot, as shown in the image.
MitoSciences immunocapture antibodies and kits can be used to isolate large enzyme complexes in their fully intact, fully active states from small samples of tissue or cells. For kits, immunocapture antibodies are irreversibly cross-linked to protein G-agarose beads, which allow for the purification of enzymes as a subsequent analysis of subunit composition, and post-translational modifications. Immunocapture antibodies are also available separately for incorporation into your own applications. An example of this is shown in the figure where ATP synthase, part of the OXPHOS pathway has been shown to be immunoprecipitated using the immunocapture kit ab109715. The identity of the protein bands where shown here, were confirmed by mass spectrometry.
If you work in stem cell research, we have a range of well-tested antibodies for embryonic stem cells, such as Oct4, Sox2 and Nanog. For germ cell research, we offer, for example, DDX4, DAZL, Stella, GCNF and PIWIL2. For cancer studies these are some of our best antibodies that we would recommend for your research. If you are studying cancer metabolism, we have mTOR, TGF beta and VEGF, amongst others. For hypoxia we recommend our Hsp90 and HIF1 alpha protein antibodies. You can find an even wider selection of western blot tested antibodies for stem cell, cancer studies and all other research areas by visiting our website.
Abcam's catalogue includes over 2,500 secondary antibodies. For western blotting, secondary antibodies are mainly coupled to enzymes such as HRP and alkaline phosphatase. We also have antibodies that allow detection on fluorescent scanners. Cross-reactivity of secondary antibodies with endogenous immunoglobulins can be a concern, especially for when the sample tissue is extremely rich in immunoglobulins. To solve this we recommend using one of our pre-adsorbed antibodies, which are available as pre-adsorbed against up to three or eight species. Should specificity of your secondary antibody be an issue, we recommend using one of our Fc-specific antibodies. In contrast to secondary antibodies, recognizing the heavy and light chain, Fc secondary antibodies only bind to the heavy chain of the primary antibody. For example, a goat anti-mouse IgG H&L antibody could potentially bind the light chains of other mouse immunoglobulins than IgG. This risk is abolished when using a goat anti-mouse IgG Fc secondary.
When using heavy and light chain-specific antibodies for western blot detection of immunoprecipitation samples, two bands can occur/appear. The heavy chain at around 50 kDa and the light chain at around 25 kDa. In the instances where your protein of interest, which has been immunoprecipitated is in the 50 kDa range, the heavy chain can obstruct the detection of the band of interest. To solve this problem, simply use one of our light chain-specific secondary antibodies, which will no longer recognize the heavy chain.
Just to remind you, you can find out additional information about the product mentioned in this overview on the link shown on the slides, as well as in a PDF copy of the webinar, which will be available for download at the end. For more hints and tips, you can find information about western blotting protocols, guides and frequently asked questions; and also our Optiblot range on our Abcam website, using the links shown.
The Abcam scientific support team will be happy to help you with any queries you may have. Support is available in a number of languages, including English, Chinese, Japanese, Spanish, German and French. For those in North or South America, you can contact us at our US office. For Hong Kong, China and Asia, you can contact our Hong Kong office. For UK and Europe, you can contact our UK office and for Japan, our Japan office.
As a thank you for viewing the webinar, we are also offering a promotional discount of 25% on the products mentioned in this webinar. To find out more about this promotion and start saving, please visit the link shown. Thank you for your time. I am now going to pass you back to Caitlin, who will answer some questions which have been received.
CB: Thanks to everyone who sent in questions. We'll go over as many as possible, time permitting. It looks like a lot of people have questions about using loading controls about the same protein, about the same size as the protein of interest. So there are a couple of different options that you have when using loading controls. So beta actin is one of the more commonly used loading controls, and that's about 38 kDa. But if your protein of interest is around 38 kDa and you still want to use beta actin, you can probe with your beta actin antibody and then strip that antibody from the membrane, and then use your other antibody. You can certainly do that. We do have a protocol for stripping and re-probing on our website, on the Abcam website and the scientific support section; there is a protocol for doing that. I would recommend just using a loading control that is at a different molecular weight.
There are several different options for loading controls for any kind of protein, or any kind of sample that you have, so it would probably be easier to just use a different loading control. The benefit of doing that, if you have a loading control that's a significantly different molecular weight than your protein of interest, you can actually, after transfer, you can cut the membrane and then stain using those two different antibodies completely separate from each other. That way you won't have to go through the extra step of stripping the membrane and re-probing. Every time you strip you can lose some of the protein off of the membrane, which can affect your results if you're trying to quantitate or just qualify the amount of your protein of interest between samples. So I would recommend choosing a different loading control than one that's close to your protein of interest.
There's a question about gradient gels from Azada; thank you, Azada. Do you have a specific protocol for preparation of gradient gels and are they available commercially? I would recommend just purchasing them. You can make them, but it would be much easier to just purchase them and they're available in different ranges of acrylamide percentage. A common one is just 4-20%, and with that range you can analyze a very large number of sized proteins. So I would recommend just purchasing a 4-20% gradient gel.
We have a question from Dev about how long do primary antibodies typically last? I used a five-year-old antibody and didn't see any signal, and could the antibody have degraded? The answer is yes. Five years, an antibody will probably be viable if stored at minus 20°C, but they can certainly degrade upon storage, so I would recommend if the antibody was working fine prior to storage, just go ahead and get a new antibody, it probably has been damaged upon storage.
We have a question from Luke about - asking for some more advice for phosphorylated proteins, to repeat again what to use for phosphorylated proteins. So the first step is make sure you include phosphatase inhibitors during your sample preparation, as well as protease inhibitors, if you aren't already doing that on a regular basis. The second step that we mentioned was to use BSA, you can use 3-5% BSA as your blocking buffer instead of the milk. So the milk has enzymes in it naturally that can break down that phosphoprotein, and then your antibody won't be able to bind to it.
We had a question from Balaji asking if blocking peptides are available for phospho-antibodies? Yes, they are. Generally, for most phospho-specific antibodies there is a peptide available. If it's an antibody that detects one specific phosphorylated residue on the protein, the immunogen would have had to be that specific region. So you can make a peptide if there's not one commercially available, but you would know that you would have to make a peptide to that region and phosphorylated at that specific residue.
There's a question from Moosu: I sometimes have spots around my bands when I develop my membrane, I have increased washing periods and based on the Ponceau there are no air bubbles. I'm assuming spots might be kind of a speckled appearance just across the membrane, and sometimes you see that it's the blocking solution, the BSA or the milk powder isn't fully dissolved in the solution, in the TBST, and so you end up with some speckles or some dots on your membrane. There could also be some contamination in one of your solutions, and that can show up as dots or speckles. So if you think it might be contamination, of course, you can either sterile filter your solutions or you can make new solutions. If you think you're having some trouble fully dissolving your milk or your BSA powder in that TBST, you can also filter your blocking solution after you add the powder. That can get rid of any speckles that might be forming, but it's not an issue with the transfer, so your Ponceau stain will look okay, but they're just showing up because of the presence of other things in your solutions.
We have a question from Mehmet: I don't have a homogenizer but I have a sonicator, can this be used to lyse my tissue? You can use the sonicator and that's perfectly fine. Something to keep in mind with the sonicator is the solution can get very warm, and so you'll need to make sure that you're not sonicating for long periods at a time. Sonicate for maybe 15-30 sec intervals, and then cool your solution in between the sonication periods, and just make sure your sample is still on ice, even while you're sonicating. So it's just very important to make sure your sample doesn't get hot, and that heat can damage your protein and then you might end up with high background on your blot as well.
We have a question from Kent: I keep seeing secondary antibodies that are called pre-adsorbed, what does that mean and do I need to use them for a western blot? The pre-adsorbtion is basically an extra pre-treatment that is done to some secondary antibodies, and it removes any molecules that might be cross-reacting with other species than the intended species. So if you have an anti-mouse secondary antibody that's been pre-adsorbed, that means it's been pre-treated to remove any anti-rat molecules or any anti-rabbit molecules, so that the secondary is specifically anti-mouse. You don't have to use this for your western blot, I think it would be more important for a western blot following an immunoprecipitation, but you should be fine to use a non-pre-adsorbed secondary antibody on your western blot.
We have a question from Ling: Someone in my lab ran a western blot and the blot was black, but the bands were white, can we fix this? The answer is yes. White bands on a black blot it's called inverse banding, and it's normally caused by using too much antibody if you're using chemiluminescence for detection. So the film is kind of quenched and overexposed, and you should be able to fix this by just diluting the primary and the secondary antibody further until the blot looks normal, and you see black bands on a white background again.
JR: Thank you, Caitlin and Mandeep. So that will conclude our webinar for now. If your question was not answered, we will have scientific support get back to you within the next 48 hr so that you can have that information, and to help you out. I just wanted to run through a couple of slides for our upcoming meetings, and we have a Stem Cells to Tissue Conference coming up in April. We have an Origins of Tissue Stem Cells, which is in Edinburgh in June, and our upcoming webinars. So we hope that you enjoyed this webinar, and we hope to see you in the future. As we mentioned before, if you do have any questions about western blots, any other scientific enquiries, our scientific support team will be very happy to help you and can be contacted at email@example.com. If you have any webinar or event-related questions, you can also contact the events team at firstname.lastname@example.org. So thank you for participating in our webinar, I'd like to thank Mandeep and Caitlin for being our presenters today. We're glad that you participated and we hope to see you again in a future webinar. Thanks very much and take care!