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Western Blot Tips and Tricks Filling the Gap Between Art and Science
Webinar July 30, 2014
[0:00:00] Slide 1 Sean Sanders: Hello everyone and a very warm welcome to the Science/AAAS webinar,
My name is Sean Sanders, Editor for Custom Publishing at Science, and I'll be the moderator for today's webinar.
In today's event, we will be investigating the art and science behind
Western blotting, a technique used in practically every life science's laboratory by thousands of scientists worldwide. Despite its popularity, the method remains as much of a skilled art as a science and is often a significant challenge to produce reproducible and reliable results.
Our three expert panelists in the studio today will give us their hints and
tips of how to get the best possible Western blot data using current techniques as well as take a glimpse into the future of how technological advances could bring the needed standardization to Western blotting.
It's my pleasure to introduce those speakers to you now. They are to my
left Prof. Pier Giorgio Righetti from the Polytechnic University of Milan in Milan, Italy; Dr. Biji T. Kurien from the University of Oklahoma Health Sciences Center in Oklahoma City; and Dr. Nick Thomas from GE Healthcare in Cardiff, Wales. A warm welcome to all of you! Thanks for being with us.
Before we get started, I have some important information for our
audience. Note that you can resize or hide any of the windows in your viewing console. The widgets at the bottom of the console control what you see. Click on these to see the speaker bios or to download a PDF of the slides.
Each of our guests will give a short presentation followed by a Q&A
session during which they will address questions submitted by our live online viewers. So if you're joining us live, start thinking about some questions now and submit them at any time by typing them into the box
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on the bottom left of your viewing console and clicking the "submit" button.
If you can't see this box, click the red Q&A widget at the bottom of the
screen. Please do remember to keep your questions short and concise. That will give them the best chance of being put to our panel.
You can also log in to your Facebook, Twitter, or LinkedIn accounts during
the webinar to post updates or send tweets about the event. Just click the relevant widgets at the bottom of the screen. For tweets, you can add the hashtag #ScienceWebinar. Finally, thank you to GE Healthcare for sponsoring today's webinar.
Before I kick off the presentations, I'd like to put up a quick poll for our
viewers. Please make your choice now and click the submit button. We'll come back to the results of the poll during the Q&A portion of the webinar. Now, I'd like to introduce our first speaker, Prof. Pier Giorgio Righetti.
Prof. Righetti is a full professor of Proteomics at the Polytechnic
University of Milan. He is on the editorial board of numerous journals and has co‐authored a 2013 book on proteomics with Dr. Egisto Boschetti.
Some of the technologies developed by Dr. Righetti include isoelectric
focusing in immobilized pH gradients, multicompartment electrolyzers with isoelectric membranes, membrane‐trapped enzyme reactors, temperature‐programmed capillary electrophoresis, and combinatorial peptide ligand libraries for detection of low‐abundance proteome.
Recently, Dr. Righetti won the 2014 HUPO Award for Distinguished
Achievements in Proteomic Sciences. A warm welcome, Prof. Righetti! Slide 3 Dr. Pier Righetti: Thank you, Sean, and thanks to all of you for being with us today. I think I
would try to tackle this topic from a perhaps unexpected point of view, which is basically that I believe that one of the major problems in Western blotting is the protein load.
Slide 4 We typically work with loads as low as 1 nanogram, even below 0.1
nanogram, which is the detection limit of silver staining. At such low loads, blots can be poorly reproducible due to antigen losses even during
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the sample preparation, losses due to adsorption onto glass and plastics, losses due to adsorption even to the polyacrylamide gel fibers, and incomplete electrophoretic transfer, of course.
Due to these accidents, it is no wonder that trace antigens have very
small chances to be detected. I think there could be a solution to it and this remedy could be sample pre‐fractionation, which is something that people often overlook, but we in proteomic science do have two major methods for reduction of the sample dynamic range.
Slide 5 Remember, the sample dynamic range in any of your sample can be as
much as ten orders of magnitude, which is really a huge, huge interval. So we have two methods, one called the Subtraction Approach by which we take a serous of 20 or even many more anti‐sera. We try to remove the most abundant proteins and I believe there are quite a few problems with that.
The other approach is the Enrichment Approach by which we try to
enrich categories like phosphoproteins, glycoproteins, and the method we have is more ecumenical.
[0:05:09] It's broader. We try to accept every protein in your sample and this
method that we developed is called combinatorial peptide ligand library. This method allows you for instance to handle very large volumes of your sample. We have even treated up to 1 liter volume.
And it acts simultaneously by reducing the high‐abundance proteins
which are pestilent in the detection of any low‐abundance species, and simultaneously enriching the low‐abundance proteins. Often we get an increment of visibility for these low‐abundance species up to three, sometimes even four orders of magnitude.
So what is the chemistry of these combinatorial peptide ligand libraries
and what is their action mechanism? We will see it in this slide. Slide 6 This chemistry, we prepare them by a combinatorial chemistry, so we
hook 20 different amino acids to these polymethacrylate beads via a
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spacer and a linker, and it's a classical Merrifield chemistry, except that it's on combinatorial.
At the end of the process, we make only six amino acid long peptide, so
we only work with hexapeptides, but these hexapeptides could be as many as a diverse, huge population, as many as 64 million diverse baits, each one to collect and hook on to a specific protein. So, how do they function? Here it is.
Slide 7 Basically, you take your sample, which has a broad, very large dynamic
range and you load it not necessarily into kernel. We can do it batch‐wise. You can take any volume and drop tiny amounts of beads. Sometimes, we work with only 100 microliters of beads. You let them equilibrate for a couple of hours then we wash out the excess of whatever that's not been bound under native conditions.
And then under denatured conditions, we collect the sample that had
been bound. So as you can see in the lower right, the dynamic range of the collected sample is much, much different from the initial sample, so you have a much higher chance to see the low‐abundance components.
Slide 8 Now, to explain better this mechanism of action, when these samples are
incubated with the beads, the high‐abundance proteins quickly saturate your specific ligands, which means that we only capture a tiny, tiny amount of the high‐abundance species. The remaining gets lost in the washing process, whereas the low‐abundance species could be enriched on a specific ligand bead. And so, you can get as much as you want provided you give to the beads larger and larger volumes of your sample. That's the way.
Slide 9 Finally, what you end up is this situation. The initial situation you can see
in the upper graph to the right. We have these huge peaks, which are the high‐abundance proteins, which obliterate the signal of the low‐abundance proteins, and the valleys are the low‐abundance components.
The valleys are enormous as compared to the peaks in polydispersity. At
the end of this process, if you're lucky, we try to bring the vast majority of our proteins within the range of detection of your, let's say, mass
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spectrometer, blotting units, Western blotting and so on, so this is the real mechanism. Now, let's see how they work.
Slide 10 Let me show you. This is a nice example. We took human serum in which
we believe that the dynamic range could be 12 orders of magnitude and we spotted it with 1 to 160 nanograms per microliter of serum amyloid A. What you see in these eight panels are tiny sections of 2D maps taken in the lower range of masses and acidic ranges. As you can see, zero nanogram is the control and then you see the different notes from 1 nanogram to 160 nanograms.
You can easily see that you begin to see something at a load of 10
nanograms. You see these red circles. 20 nanograms are visible and so on, but below 10 nanograms, we essentially don't see anything.
Slide 11 Now, the same one, the same serum treated with these combinatory
ligand libraries, look at that. I want you to look at the second panel, 1 nanogram. You see already how visible are these spots.
The second spot is so dense that in reality ‐‐ let's go back ‐‐ Slide 10 It's as dense as the 80 nanogram load, which means that basically ‐‐ Slide 11 And then you see a constellation of spots all around because you also
enhance the visibility of other low‐abundance species. So you can see that by this technique, we enhance the visibility of your low‐abundance species by perhaps close to two orders of magnitude.
[0:10:09] Now, let us go to another real sample. Slide 12 This is a sample trying to detect allergens in maize kernels. Maize has
been introduced to Europe 500 years ago and yet up to today, only one
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allergen had been identified, which is the lipid transfer protein, which is ubiquitous. You can find it anywhere. We thought it is impossible that it is just a single allergen, so we treated this maize extract with our combinatorial ligand libraries.
As you can see from these three other panels, which are blot to the right
with these red circles, we could identify many more allergens that had been invisible before and we identified five more. So we went from one to six ‐‐ Vicilin, globulin‐2, endo‐chitinase, thioredoxin and trypsin inhibitor. And here on the left, you can see the Coomassie stain, SDS monodimensional profile, and different blots for different patients. This is just one example.
Slide 13 Now, let me show you ‐‐ we did recently some tropical fruits and it is
interesting. For instance, in avocado, only one antigen had been known, Pers a 1. Here you can see the blots, immunoblots from 16 different patients extracted in two different conditions, either phosphate buffered saline or TUC, which is thiourea, urea and CHAPS. You can see the bands of the reacted allergenic proteins.
After this combinatorial ligand treatment, we could detect not just one,
this Pers a 1, but five more allergens ‐‐ profilin, a polygalacturonase, a thaumatin‐like protein, a glucanase, and an isoflavone reductase‐like protein. You can see a big jump from one to six, one to seven and so on, so it really shows that this technique has much to offer in terms of detecting antigens and allergens in your sample.
Slide 14 This is another example, a banana. In a banana, there were five allergens
already reported, which are called Mus a 1, Mus a 2, Mus a 3, Mus a 4, and Mus a 5. Of course, we detected all of them, but additionally after the combinatory ligand library treatment, we could detect another two allergens that have been previously unreported, a pectinesterase and a superoxide dismutase.
I would also like you to notice that as much as we enhance the visibility of
allergens, we also enhance the visibility of plenty of other low‐abundance proteins that had never been reported up to present time.
So for instance, the control, untreated one, this violet bar that you can
see, we could only detect 452 unique gene products, unique proteins. But
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after this treatment, we could detect 849, so about twice as many as compared to the control. That tells you the power of this technique in detecting everything, not just allergens and antigens, but any other low‐abundance species.
Slide 15 Now, the last example I want to show you is this mango. Mango, as you
know, is a tropical fruit and it is used a lot in the kitchens in Eastern countries and so on.
In mango, very little was known. There were essentially no reports, no
studies on that. Again, what I want to show you in the control, we took the pulp. We homogenized it. We extracted it. In the control, untreated sample, we could detect only 374 proteins, but after the treatment with this combinatorial peptide ligand library, we could detect close to 3000 proteins. So here, we have an increment of visibility of almost one order of magnitude, which is quite unique.
And then in terms of allergens, no allergen had been reported so far, but
after this treatment, we could detect of course the lipid transfer protein, which is as I told you a universal allergen found in any type of food, essentially superoxide dismutase, germin‐like protein and profilin, so we could detect at least four allergens. Maybe there are more that have escaped our dictation, but still it's a big step forward.
I would like to finish my brief presentation by telling you that sometimes,
you have the feeling especially in this field of Western blotting where so many problems are reported and will be treated by the other two speakers.
[0:15:01] Slide 16 Sometimes or even most of the time, you have the feeling that your prey
is escaping capture, and here is a nice presentation. This pig is trying to escape that sentence. And so, if you don't capture your pig, you will end up with an empty dish. You will have to skip your meal, which is what happens most often.
Slide 17
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Now, maybe we have a remedy for you. This is the cover of the book that we wrote together with Egisto Boschetti. This book deals very, very extensively with this topic and it has ‐‐ at the end of it, there are 50 pages of protocols, so any protocol, steps and procedures, including some protocols in Western blotting, so maybe this will be your lifesaver. Thank you so much for your attention. Thank you.
Sean Sanders: Great! Thank you so much, Prof. Righetti. Slide 18 Our second speaker for today is going to be Dr. Biji Kurien. Dr. Kurien's
research interests include the study of free radical, mediated damage in systemic lupus erythematosus and Sjögren's syndrome as well as the role of the nutraceutical curcumin in autoimmune diseases.
His publication record includes numerous publications in national and
international peer‐reviewed journals. In addition, he co‐edited two volumes entitled "Protein Blotting and Detection and Protein Electrophoresis" as part of the Methods in Molecular Biology series.
Welcome to you, Dr. Kurien. Slide 19 Dr. Biji Kurien: Thank you, Sean. It's a privilege to be here today for this Science
Webinar, as well as to participate alongside Dr. Righetti and Dr. Thomas. I'm grateful to you, AAAS and GE, for the invitation to present on tips and challenges in Western blotting.
Slide 20 Western blotting involves the transfer of protein patterns from gel to
microporous membranes. Electrophoretic transfer was first described by Dr. Harry Towbin and then later by Dr. Neal Burnette. Dr. Burnette coined the term Western blotting to retain the geographic naming tradition initiated by Dr. Southern's paper.
Slide 21 There are several advantages of transfer to membrane. Wet membranes
are pliable and are easy to handle. This allows easy accessibility of proteins immobilized on membrane to different ligands. Only small amount of reagents are required for transfer analysis.
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Slide 22 It is possible to obtain multiple replicas of a gel. This allows for prolonged
storage of transferred patterns. The same protein transfer can be used for multiple successive analyses.
Slide 23 There are three kinds of Westerns ‐‐ the good, the bad, and the ugly. Slide 24 One can take several steps to obtain a good Western blot. This includes
proper sample preparation, the choice of right lysis buffer, proper gel preparation, right electrophoresis and transfer conditions.
Slide 25 When one prepares samples from cultured cells, it is customary to wash
the cells with PBS to remove cell culture media. It is important to remove PBS completely prior to cell lysis. This is because salt contamination can cause horizontal streaking in two‐dimensional gel electrophoresis.
Slide 26 To eliminate viscosity from DNA in crude cell extracts, it is essential to
treat cell extracts with protease‐free nuclease. This can also be accomplished by sonication or vigorous vortexing of heated sample.
Slide 27 There are several lysis buffers to solubilize proteins. Laemmli lysis buffer
has been used to solubilize cell extracts for SDS‐PAGE. Urea/thiourea containing lysis buffer has been used to solubilize proteins for two‐dimensional gel electrophoresis. This buffer has been especially found to be useful to solubilize small HR proteins. Radio‐Immune Precipitation Assay buffer has been found to be useful to solubilize smooth muscle proteins. One can also personalize lysis buffer to solubilize a protein of one's own interest.
Slide 28
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SDS to protein ratio, 1.4 microgram of SDS binds to 1 microgram of most proteins. It is better to have excess SDS compared to protein sample. Dr. Hames recommends SDS to protein ratio of 3:1. Therefore, it is important to determine protein concentration to prevent inadequate protein to sample buffer ratios.
Slide 29 SDS does not unfold certain proteases. Therefore, it's essential to heat
sample soon after adding lysis buffer. This avoids protein degradation by proteases. A very small amount of protease can degrade proteins if the sample is not heated soon after adding lysis buffer.
[0:20:06] Slide 30 Do not heat sample with lysis buffer at 100 degrees centigrade for more
than five minutes. This is because the aspartyl‐proline bond is most susceptible for cleavage by heat and acidic conditions. Heating it at 75 degrees centigrade for five minutes avoid bond cleavage. It also helps to inactive proteases.
Slide 31 Heating with SDS lysis buffer alone may not solubilize proteins or
membrane proteins. One may need to add 6 to 8 mole urea or Triton X‐100 for this purpose. Insoluble material should be removed by a high speed spin for a couple of minutes. This will help avoid streaking within gel.
Slide 32 Protein aggregates can form in SDS samples. This is because reductant
becomes partly oxidized upon cooling sample, letting part of the cysteines unprotected. This can lead to back‐folding and creation of inter‐polypeptide aggregates which can in turn lead to blurred zones and formation of double bands.
Slide 33 Cooling sample of about 60 degrees centigrade and adding an alkylating
agent avoids this problem, which can lead to sharper bands and abolishment of artifacts.
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Slide 34 Miscalculating cross‐linking factor. The pore size of a polyacrylamide gel
is dependent on two factors, (a) the total concentration of acrylamide referred to as "T", and (b) the degree of cross‐linking referred to as "C". We mistakenly assume that given total acrylamide concentration T is the percentage of acrylamide per volume, and C is the percentage of bis acrylamide per volume.
Slide 35 To avoid this, one can calculate percent T or percent C by the following
formulas. T = [(a+b) x 100] / v; C = (bx100) / (a+b) where "a" is equal to the mass of acrylamide in grams, "b" is equal to the mass of bis acrylamide in grams, and "v" is equal to the volume. Alternatively, one can use commercially available acrylamide/bis acrylamide solutions for making gels.
Slide 36 Overloading and underloading are the two commonly encountered
problems in electrophoresis. Overloading leads to distorted bands in lane, as well as in adjacent lanes. Underloading leads to poor detection of minor bands.
Slide 37 It is not necessary to adjust the pH of the running buffer in SDS‐PAGE
even though the pH is above 9. Adjusting pH to 8.3 leads to higher load of chloride, which leads to longer protein separation times and some zones remain poorly resolved.
Slide 38 It is best to polymerize SDS‐PAGE overnight at room temperature to
ensure complete polymerization. However, Dr. Haeberle described a gel with greatly accelerated rate of polyacrylamide cross‐linking. One can run this gel in a dramatic fashion in five minutes using the Haeberle running buffer heated at 70 degrees centigrade.
Slide 39
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It takes about two to four hours to run a standard mini‐gel. We have found that it is possible to run a pre‐cast gel in ten minutes using heated Laemmli running buffer. The next couple of slides will show very recent results, actually, the results we got about the day before yesterday evening showing that electrophoresis, Western blot, and immunoblotting can be accomplished in a few hours with actual run time of 60 minutes.
Slide 40 This slide shows the Western blot of protein marker. The pristine protein
molecular weight marker was electrophoresed on a 4% to 20% precast gel in ten minutes and transferred to nitrocellulose membrane in seven minutes using a semi‐dry transfer system. Lanes one to four shows the molecular weight markers.
Slide 41 This slide shows the result of an ultra‐fast bovine La immunoblot. Bovine
La, purified bovine La was purchased from ImmunoVision and this antigen was electrophoresed on a 4% to 20% pre‐cast gradient gel in ten minutes and transferred to membrane in seven minutes.
The subsequent immunoblotting steps were all carried out at 37 degrees
centigrade. All the reagents were pre‐warmed to 37 degrees centigrade prior to adhesion to the membrane. The membrane was blocked with milk for ten minutes and then incubated with a primary antibody for ten minutes. The membrane was then washed three times with deionized water and three times with TBST all in three minutes.
[0:25:05] Then the membrane was incubated with secondary antibody for ten
minutes and washed as described previously for three minutes. Then the NBT/BCIP mixture was added and the bands appeared in less than 30 seconds.
Another antigen was experimented similarly and the results obtained
were only the positive control bound; the negative and the conjugate control did not bind. This result shows that the entire process of electrophoresis, Western blot, and immunoblot can be carried out in a very rapid manner.
Slide 42
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There are two types of transfer buffers, Towbin transfer buffer and the CAPS buffer system. Towbin buffer system contains Tris, glycine, methanol, none to 0.01% SDS. CAPS buffer system contains methanol, CAPS at a pH of 11. This buffer is used to transfer proteins to PVDF prior to in situ blot sequencing.
Slide 43 Transfer buffers without SDS are better. This is because proteins can pass
through the PVDF membrane in the presence of SDS. However, SDS is required during transfer of proteins that have a tendency to precipitate.
Slide 44 Methanol in the transfer buffer helps in stripping SDS from proteins. It
stabilizes the geometry of the gel during transfer. It increases the binding capacity of nitrocellulose from protein and helps proteins to bind better to nitrocellulose.
Slide 45 Transfer buffer without methanol is useful to transfer high molecular
weight proteins, for transferring conformation sensitive antibodies, and when enzyme activity needs to be preserved. However, the drawback is that PAGE gels tend to swell in low ionic buffers. The bands tend to get distorted if the swelling occurs during transfer.
Slide 46 When blotting proteins from SDS‐PAGE, the proteins are eluted as
anions. Therefore, the membrane should be placed on the anode side of the gel. It is important to remove bubbles trapped between nitrocellulose and gel to prevent bald spots. The gel plus filter plus pads assembly should be tightly held together to ensure good transfer and no band distortion.
Slide 47 Do not exceed the binding capacity of membrane because excess protein
can get weakly associated with membrane and is readily accessible to antibody. However, the ensuing protein‐antibody complex will wash off in the subsequent immunoblotting steps leading to reduced signal.
Slide 48
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High molecular weight proteins transfer very poorly. This can be avoided
by using reversible gel cross‐linkers followed by gel depolymerization prior to Western blot transfer, limited protease digestion during transfer, prolonged transfer with addition of SDS, and heat mediated transfer in tank buffer systems.
Slide 49 Low molecular weight proteins tend to bind with low affinities to
membrane. Such proteins can be lost during transfer or washing. This can be avoided by cross‐linking proteins to membrane, and the use of 0.2 micrometer membrane can reduce the phenomenon of "blow through" through which these proteins can be lost.
Slide 50 To conclude, Western blotting is a simple method. The potential of
Western blotting has greatly evolved. There are huge numbers of ways to transfer proteins. This method has led to a deeper understanding of protein ligand interaction. This concludes my presentation. Thanks for listening.
Sean Sanders: Thank you so much, Dr. Kurien. Just a reminder to those watching the
webinar that I know there was a lot of information on those slides. Thanks, Dr. Kurien, for putting all of that together, a lot of great advice. If you want to download a PDF of the slides, just click the green widget at the bottom of your screen labeled "resources list" and you can get a PDF right there.
Slide 51 We're going to move on to our final speaker now. That's Dr. Nick Thomas.
Dr. Thomas is currently a principal scientist in Life Sciences at GE Healthcare based in Cardiff, Wales. In a 30‐year career in industry, he has held a number of positions in Operations, Marketing, and Research and Development.
He is the inventor or co‐inventor of over 60 patents covering a wide
range of technologies including microfabrication, molecular and cellular sensors, and cellular imaging. Dr. Thomas received the GE Edison Award for contributions to research and development in 2012 and is currently an honorary visiting professor at Cardiff University.
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Slide 52 Welcome, Dr. Thomas. Thanks for being here. Dr. Nick Thomas: Thanks, Sean. It's great to be here. So our previous two speakers have
given you some great technical tips and information about how to get the best out of your Western blotting. I'd like to spend a few minutes giving you my perspective of 30 odd years of doing Western blotting in various guises and where I think the technique is going in the future.
[0:30:08] It seems somewhat strange to me to think back to when I started
Western blotting. At that time, blotting was a standard method for detecting not just proteins, but also DNA and RNA and Southern and Northern blots. Of course, PCR has killed all of that. So I guess until somebody gets a Nobel Prize for working out how to amplify proteins, we're still going to be doing Western blotting for some time.
Slide 53 As Sean said in the introduction to the webinar, Western blotting is a very
widely used technique in many labs worldwide and there's a huge amount of information and data published every day on the technique. It's still a key technique for looking at protein expression, characterizing proteins, and looking at post‐translational modifications like phosphorylation.
But despite its widespread use around the world in different labs, all of
those different labs tend to use different workflows. Even different people within the same lab will use different workflows and that has potential to have quite significant variability in data, and the technique is highly dependent on skill and experience, and probably about one in four, one in five Western blotting experiments don't get too good results.
Slide 54 So there's obviously, I think, need for improvement. One thing is to get
around common errors that people make or things that happen during the course of their experiments. I've put together a row of gallery on the right hand side of this and it shows some of the common things that happen when people are doing their Western blotting experiments when they're in a hurry to get data for publication because the lab boss is screaming at them for that data.
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Everybody will recognize things on this, I guess, that's happened to them,
things like the one in the top left there. You've run your gel. You've cast your gel or you've run your gel. You've got a nice gel then you go on to put your blotting stack, so you very carefully put your blot and gel together, get all the bubbles out. You don't crack the gels. You put it in the blotter and then you connect the stack the wrong way around and you wave goodbye to your sample.
As I said, I think there are areas that need improving in Western blotting,
standardizing, improving the reproducibility of the process, getting better quantitation, simplifying the workflow, basically to get it right every time and basically to free up research time.
Slide 55 So in terms of my experience, as I said, I started a long time ago. In those
days, we were using colourimetric detection, DAB‐HRP detection on nitrocellulose. At that time, I was and still am a very keen photographer. We published a number of methods adapting photographic processing techniques to Western blot processing, improving the background, reducing reagent usage.
At that time, my group also developed the first pre‐stained molecular
weight markers for SDS‐PAGE in Western blotting and we worked on the first commercial ECL detection system.
Slide 56 I mentioned I'm a keen photographer. This is Ansel Adams, an American
photographer and in my opinion, and I guess in the opinion of many people, the greatest landscape photographer that's ever been. The key thing that made Ansel Adams so well‐revered is basically summarized by this quote from him. "You don't take a photograph; you make it."
Here's a picture of him taken in the 1940s with a plate camera about to
expose. Basically, his photography was not about taking photos just by pressing the shutter button. He had a very standardized, rigorous process from visualizing the image that he wanted, going to the location, exposing his film correctly, exposing the prints correctly, a very complex, standard way of making prints, which I think has a lot of commonality with Western blotting.
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Western blotting, I think, starts with the time at which you visualize your experiment. You visualize what type of data it is. You plan your experiment. You go through the process and then you interpret your data. And the real skill in Western blotting, I think, the science in it is that at the beginning of the process, visualizing and planning your experiment, and at the end of the process, interpreting the data. That's where the skill comes. The bit in the middle, really it's cookery in my opinion.
Slide 57 So let's look at the parallels between the evolution of photography and
the evolution of Western blotting. This slide illustrates basically what I was doing when I first started processing black and white prints 30 odd years ago. You start with film. You go through a whole series of operations, which are basically sloshing stuff around in chemicals. I can see the similarity to Western blotting there and you end up with a print.
[0:34:59] I still got a lot of the contact sheets and the prints that I made in those
days in filing cabinets. I still get them out and look at them occasionally, but the processing trays that I used to develop those contact prints have now no use and they now in fact make very good snow shovels.
Slide 58 Nowadays, like many photographers, I've converted entirely to digital, so
I can go from a digital raw negative in a camera on location, take that raw negative into a computer, process it, using many of the same techniques that I used for wet chemistry in developing. I can change the contrast.
I can selectively dodge and burn different areas, but the key thing is what
digital gives you, it frees you from that tyranny of the dark room. You can establish a reproducible workflow. As illustrated here, I can go from Adobe light room straight into a final process print, which then gets sent off to a publisher for a magazine cover.
Slide 59 That evolution, I think, is very similar in many ways to the evolution of
Western blotting from 20th Century techniques if you like to 21st Century techniques.
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We started, as I mentioned, colourimetric detection using either HRP or alkaline phosphatase on nitrocellulose membranes, then we moved on to chemiluminescent detections, better membranes on PVDF, and the advent of imagers and scanners and so on replacing in a lot of case X‐ray film and it brings much better quantification, which is really what it's all about. That's really in the end what a Western blot is to do, is to measure. Okay, something is present, but it's much better to know quantitatively home much is there.
And if we move into the more recent history, moving from film to
detection to digital imaging, using fluorescence labels to do Western blotting, and that's brought out a lot of techniques which basically when I started weren't dreamed of, the ability to do multiplexing in two or three colors to ask much more complicated biological questions.
Slide 60 If we just touch on a couple of technical aspects, here I'm showing you
improvements in dynamic range using solid state CCD detectors against X‐ray film, so you get a much wider dynamic range with CCD imaging. And as I mentioned before, using fluorescence has a great advantage of doing multiplexing.
In this example shown on the right, you can look at the amount of ERK,
but you can also use a housekeeping protein such as GAPDH to normalize within your bands.
Slide 61 You can take that a stage further. Showing here on the left hand side, the
measurement of ERK, phosphor work [Phonetic] and a housekeeping protein. As I said before, that type of experiment gives you very good quantification of the involvement of a protein and a signaling pathway within a cell that you couldn't have dreamed of or even attempted to do using standard techniques that we had years ago.
Slide 62 So in terms of Western blotting evolution, I think it's really poised for
evolving to its next stage and that will be taking all the types of things, the sample preparation, the electrophoresis, the blotting, everything that's shown on the right hand side there, and going through a process of systems integration basically aiming to get protocols that are standardized, quantitative and smart and free up science.
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Slide 63 Basically, just to conclude, I think that systems integration, which is
coming very quickly I think now, streamlining the workflow, removing those errors, those opportunities for errors that I showed you, still providing light digital photography, the artistry. You still need the same skills. Now, it's an often used maxim that digital cameras or better cameras don't make you a better photographer, and that's exemplified by all the rather bad images that you can see on the web these days from camera phones.
So basically, systemizing but basically allowing the skills of a scientist still
to be applied at the important place, which is at the beginning, I think, in the design of the experiment and in the interpretation, which is all about better science in the end.
Slide 64 I think all of that will come into a systemized workflow and as a
byproduct, you'll have a better place for your lab goldfish to hang out. Slide 65 So with that, I'll finish. Thank you for your attention. Sean Sanders: Great! Thanks so much, Dr. Thomas, and many thanks to all of our
speakers for the great presentations. Before we move on to questions submitted by our online viewers, I'm
going to give everyone a final chance to answer our poll, which should be up in your slide viewer right now. Give me one second. I think it should be coming up for you.
[0:40:01] So just click on the answer that you would like to choose and hit the
"submit" button and it should get through to us. We'll look at those poll results in a minute.
For those watching us live, you can still ask questions by typing them into
the text box and hitting the "submit" button. If you don't see the text box on your screen, just click the red Q&A icon and it should appear. Let's take a look at our results. I'm going to send those out to the audience.
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Well, it seems that about 26% of our audiences think it's simply an art,
18% is science, 8% a burden, and 46% all of the above. I guess I would expect something like that. I think we all agree that it's a bit of an art and a science, but it seems that for a lot of people, it's also a burden.
Maybe I can ask each of you very quickly to talk about when you're
looking at a Western blot protocol, what do you think are the most critical steps for somebody either starting a Western developing experiment or who's had some experience should be looking at? Dr. Righetti?
Dr. Pier Righetti: I think that just about all the steps are critical in a way, but I repeat, my
personal view is that if you don't enhance somehow the concentration of your highly diluted antigen by any technique, of course, we suggest the combinatorial ligand library, but often you have to concentrate it somehow by a factor of one to two orders of magnitude. Then your Western blot likely will succeed. That's my personal feeling.
Sean Sanders: Dr. Kurien? Dr. Biji Kurien: I believe that every step is critical for getting a good Western blot result.
However, if I was to pick two important aspects, I would say the position of the membrane during the transfer, if you mess up that, you lose all the proteins. Finally, if you add the wrong conjugate, then you get nothing.
Sean Sanders: No signal, right. Dr. Thomas? Dr. Nick Thomas: They're all important in that if you make a mistake in any of them, you'll
ruin your experiment. I think my view is that the very first step is the key one, and that's the design of your experiment. I think that's where a lot of the scientific skill comes in, and also at the end in the interpretation.
The other thing I would say is yes, all the processing steps, the detection
steps, the best thing to do is to standardize those and to use the best techniques available for each step, and then have the effective technique.
Slide 66 Sean Sanders: Excellent! Great advice. I'm going to get on to some more specific
questions that we've had come in. Let me start maybe with you, Dr. Kurien. I've had a number of questions about handling large proteins. I
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know that it was something that you mentioned. Do you have any advice for people working with proteins 250 to 300 kilodaltons?
Dr. Biji Kurien: Yes. Dr. Greaser has published a paper using agarose SDS, SDS agarose
gel, a vertical gel, and he uses a reducing buffer in the upper chamber. This helps to run the gel like he uses ‐‐ he ran a protein titin that's over 3000 kilodalton, up to 3700. He was able to transfer that very efficiently.
The heat‐mediated transfer protocol that we published that we can
transfer in tank buffer systems, it depends upon the gel thickness. We can transfer at very varying time periods ranging from 10 to 20 minutes. It shows that you can transfer very high molecular weight proteins very efficiently.
Sean Sanders: Now, how about very small proteins? Dr. Biji Kurien: Very small proteins, we have to be very more careful because low
molecular weight proteins tend to come out first. The best way would be after you transfer, you can fix the membrane, immobilize the protein on the membrane, as well as using 0.2 micrometer membrane can help.
Sean Sanders: Okay. Dr. Pier Righetti: Can I add something about the high molecular mass proteins? There is a
technique that people often overlook because it's a little bit more laborious, but basically you could cast your own polyacrylamide gel to as low as 4% T total monomers by increasing the percent C, the percent of crosslink fluorescence to 10% and so on.
[0:45:17] The gel, you enlarge the pore size substantially, but the gel can still be
handled because of the higher percent of crosslink. By the way, increasing the percent of crosslinkage also increases the pore sizes. Obviously, that requires more skills and obviously, you would not be able to use pre‐casted gels. You will have to do your own gels and that might add to the burden, but this is a technique we used a lot in the past and it's quite successful.
Sean Sanders: Great! Maybe that brings us nicely to the discussion of cast your own gels
or your pre‐cast gels. I remember my experience casting gels. You mess that up and your whole experiment is gone.
Dr. Pier Righetti: Yeah. Right.
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Sean Sanders: Dr. Kurien, I know you had some advice on that in your talk. Dr. Biji Kurien: Yeah. I would prefer pre‐cast gels to bring constant results. The variability
would be less. It's convenient also because acrylamide is a neurotoxin. You can avoid handling acrylamide.
Sean Sanders: Dr. Thomas was commenting last night that we might be banning that
one day. Dr. Nick Thomas: Well, we're not going to be around forever. Dr. Pier Righetti: It could be coming soon. It could be coming soon. Actually, at least in
Europe, they are pushing the legislation to total elimination. In fact, we are desperately trying to figure out what can substitute acrylamide, which is not easy.
For instance, you can use agarose. But immobilized pH gradient gels,
which is a classic, a first dimension of 2D maps, there is no way to cast, to graph immobilized chemicals to agarose. We've tried that for decades and we never succeeded, so an acrylamide‐free life is difficult to me.
Dr. Nick Thomas: Yeah, I would agree. Dr. Pier Righetti: What would be a substitute? We do not know. Dr. Nick Thomas: Yeah, but pre‐cast gels ‐‐ unless you have a rare example of needing a
very low porosity get that isn't within the standard ‐‐ Dr. Pier Righetti: Could I add another comment that we have been discussing with you
here last summer and so on? You could have pre‐cast gels, which are really toxin‐free because no matter how hard you try, when you cast a polyacrylamide gel, there are always given amounts, up to 5% or more, of free monomers that have not been incorporated and the toxicity comes from the double bond of the free monomers obviously. Do you understand?
So you never get rid of it, but we now have tricks to supply acrylamide‐
free, monomer‐free gels because we can treat them with scavengers so that at least if you dispose of them, there would be no toxicity for the environment, which is also another interesting point.
Sean Sanders: So I'm going to come to you, Dr. Thomas, with I think one of the more
difficult questions about normalization, so I'll throw this your way.
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Dr. Nick Thomas: Okay. Sean Sanders: This viewer asks, "What about normalization? What is the difference
between target protein normalization and total protein normalization? Is normalization important?"
Dr. Nick Thomas: Yes, normalization is important. Nobody would dream of doing an RT‐PCR
experiment without having the right normalization. Normalization, people will use ‐‐ there's a lot of debate in that community whether you use something like beta‐actin, whether you use what people would call a housekeeping gene like GAPDH.
So two issues, I guess, one is to have ‐‐ your normalization protein should
be around the same abundance as the thing you're detecting. So if you're looking for something, which is very low‐abundance, then you shouldn't really be using beta‐actin.
The other issue is if you're using a housekeeping protein ‐‐ and GAPDH is
probably the one most people use or is very commonly used ‐‐ it's clear from RT‐PCR experiments that that is not an invariant protein and particularly if you're interested in working in stem cell differentiation. GAPDH values can go up and down considerably.
So basically think about your experiment and always plan it as if you were
doing an RT‐PCR experiment, and I think that's a good analogy for lots of Western blotting. People would not dream of doing RT‐PCR with buffers that they thought might be contaminated. Well, why would you tolerate that in Western blotting?
So coming back to the normalization and I think that's a great potential
advantage of fluorescent detection in that you've got three colors, so you could if you like, if you want to do a very critical quantitation in normalization, you could have one color for your protein of interest and use two colors for two different normalization proteins.
[0:50:11] You might, for example, use beta‐actin and GAPDH, but you would
absolutely be looking to see that the ratio of those two proteins stay the same across your sample because if you didn't, then one of them is obviously changing and therefore, you can't normalize against a changing target.
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Coming back to what I was saying before, design your experiment appropriately at the beginning and think about what's going to go on or what could possibly vary in your sample. Otherwise, you're going to have to do it again.
Sean Sanders: Right. Dr. Kurien, I'm going to come to you with I guess a related
question, and that's if somebody wants to do quantitation, what is needed in order to make a gel quantitative?
Dr. Biji Kurien: So just minimize the variability from gel to gel. We have seen in diffusion‐
mediated non‐electrophoretic transfer, you can quantitate actually. It has been found that 10% of the proteins are transferred in three minutes in a diffusion‐mediated non‐electrophoretic transfer and 30% in 30 minutes.
But the major problem is for electrophoretic transfer, you have to decide
which protein you need to do because it transfers differentially to membranes. It's like a biased transfer. So it all depends upon what protein you are interested in and then work on that particular protein to adjust the conditions and reduce variability. There are several good papers to show how you can quantitate proteins. Dr. Gad Baneth has published one on how to quantify proteins.
Sean Sanders: Good! Prof. Righetti, I'm going to come to you with a couple of questions
specific to your technique that came in, the CPLL technique. First, you were asked, "Can you use this enrichment technique if you have a very small amount of starting material?"
Dr. Pier Righetti: Yes, you can. You can. In that case, you can reduce the amount of beads
that you use. For instance, when working with cerebral spinal fluid, people told us that often you cannot draw from a patient more than half a milliliter or something, so they said, "What do we do?"
In that case, you reduce dramatically the amount of beads that you use.
Normally for each experiment, we use hundred microliters of beads, but in that particular case, we went to down only ten microliters of beads and it was very efficiently ‐‐ but you have to remember that of course, that tiny sample volume, you have a tiny amount of your antigen and you collect what is in there. You can collect even 100%, but you cannot collect more than that.
So if it is still too low, that's it. You will not see it. To see it, you would
have to use a larger volume, so this is the problem. You can do it, but with limitations.
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Sean Sanders: And how would you know that your low‐abundance proteins aren't binding to the beads and remaining there?
Dr. Pier Righetti: That's a very good question. In reality, our beads ‐‐ you know, I come
from a Catholic country, so we try to be ecumenic. That's what our pope tell us, to embrace every faith, every religion, everything, but in reality, our beads are not as ecumenical as we would like them to be.
No matter how much we have now optimized everything, we still lose
some 10% of the initial sample. We do not know what's the reason for that. So basically, ideally you would want to look at the weight from your beads, but just as a control, look also at what you had in your original sample, and then combine both data.
Sean Sanders: Okay. Excellent! Dr. Kurien, I'll come back with a question for you. There
have been a number of questions in different ways about aberrations in gels, things like dumbbell‐shaped band, smiling gels, that sort of thing. The other big one is a protein that doesn't seem to match up with the size that you're expecting, so you get a nice band, but it's at a different size. So can you talk about some of the causes for these kinds of aberrations?
Dr. Biji Kurien: Posttranslational modification can lead to different sizes of protein. [0:55:05] The uneven heating of gel can cause the dumbbell type bands. We're
discussing, as Dr. Righetti was saying, the pore size. The pores can be blocked and ‐‐
Dr. Pier Righetti: Particulate material in your sample, yeah. Dr. Biji Kurien: And that can cause aberrant ‐‐ the migration of proteins. Dr. Pier Righetti: Right. Yeah. In fact, may I add, it is very important prior to loading your
sample to a gel, even if it is in SDS or so on, always give a sharp, quick, efficient centrifugation because there might be particulate material which is so dispersed, so minute that you don't see it. But when you load it, it clogs the gel pores and that is a disaster, so a quick centrifugation step will surely help.
May I also add something more? That we never load anything into two
outerlays because at the edges of the gel, the field strand is not constant as inside, so we always the two extreme empty because we always get
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curved, hearts, bands and so on due to uneven field strand distribution, field ‐‐
Dr. Nick Thomas: Put sample buffer in them. Sean Sanders: Okay, so ‐‐ Dr. Pier Righetti: Yeah. Correct. Dr. Biji Kurien: May I add one more? Sean Sanders: Sure. Dr. Biji Kurien: The heat that's running the samples using a heated buffer seems to be
avoiding all those things because the uniform heating of the gel avoids smiling completely. It runs really nice and straight. Sometimes, I see a figure on a paper where you see the dumbbell kind of thing all across the lane. It looks like maybe streaking.
But I think for uniform running ‐‐ if you're running regular gel, running
slowly and with proper sample preparation, then running it normally as without increasing the current, so you can get good bands and avoid the dumbbell thing.
Sean Sanders: Great! We're coming to the end of the webinar. We've got a couple of
minutes, so I'm just going to give you a couple more questions. Maybe Dr. Thomas, I'll come to you with this one. What is the most critical step to avoid or minimize non‐specific antibody binding?
Dr. Nick Thomas: Use a good blotter. Use a blocker. If you're talking about your primary
antibody, use a good primary antibody and use a primary antibody that's good for Western blotting. Not all antibodies to the same protein are equal. You can look at the manufacturers or the provider specification. You can believe that.
Basically, use a good antibody that you know works. Use a good
detection system that works. Use a good blocking solution, a good blocking protocol that you know that works. If you know all those things work, then you've got the basis for good protocol.
But if you've got something that works half the time and not the other
half the time, and then you've got another step which only works a quarter of the time, when you add all these things up, that's where
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people get really frustrated with the Western blotting and tear their hair out.
That's what I said before. It doesn't cease to amaze me really that people
will take infinite care in an RT‐PCR. They get their primer design exactly right. They get all their buffers clean and nuclease‐free because they know that if they don't do that, it's not going to work.
They'll take a blocking solution or a washing solution out of the fridge
that somebody else has used and they have no idea really how much bacteria are in it. They waft their membranes around, they crack them, they crease them, and they wonder why they've got marks on their films when they put it under the detector.
Sean Sanders: Good! I'm going to come with one last question to the whole panel.
Before that, actually, I want to just come back very quickly to you, Dr. Kurien. Somebody was asking if you could repeat the reference for the protein quantitation. You mentioned ‐‐
Dr. Biji Kurien: It's in the protein blotting book. I forgot the author's name. She ran the
non‐electrophoretic transfer on support, on a plastic support. She did label the protein, radiolabeled the protein and looked at the counts.
[1:00:10] Sean Sanders: Okay. Maybe we can put that up later for our viewers. Dr. Thomas, do
you have anything to add for the quantitation issue? Dr. Nick Thomas: Not particularly. Well, only to reiterate what I said before, when Western
blotting started, it was pretty much qualitative. You might have made some judgment as to there was more of protein whatever in this sample than the other.
It's very much moved to a quantitative science, but without the
development or the adoption of the best practice in the various stages, which would justify it, in my opinion, being a full quantitative science. It is in some aspects and in some labs, but I think as a technique in average, it's ‐‐ to do another analogy, it's more like PCR than it is like RT‐PCR.
You're basically looking for a band and saying, "Well, this band is darker
than that band." As I discussed in my talk, the next evolution will be in having systemized approaches using the best technologies for each step, and in some cases, bringing those altogether in an automated system.
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The whole point is doing it faster, more reproducible, and freeing up scientists that do science, not slosh things around in chemicals.
Sean Sanders: Right. Dr. Kurien? Dr. Biji Kurien: The doctor's name is Dr. Gad Baneth. Sean Sanders: Baneth, great! Dr. Biji Kurien: Yeah, for the quantification. Sean Sanders: Okay. Excellent! Slide 67 Well, I'm afraid we're going to have to end there because we are
unfortunately out of time for today. So on behalf of myself and our viewing audience, I want to thank our speakers very much for being with us today, Prof. Pier Giorgio Righetti from the Polytechnic University of Milan, Dr. Biji T. Kurien from the University of Oklahoma Health Sciences Center, and Dr. Nick Thomas from GE Healthcare.
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