Your Western blots deceive you!

Ever since the technique to blot protein samples on nitrocellulose membrane was developed, Western blot has been a go-to technique for every molecular biologist. The method is simple enough and the results seem straightforward, but correct interpretation of your results is a crucial and often overlooked step.

In Figure 1, I present an example of a straightforward Western blot result. For this experiment, I differentiated the myeloid cell line PLB into functioning granulocytes by stimulating them with dimethylformamide (DMF) for 6 days . Afterwards, I took a sample of my stimulated and control cells as well as a sample of primary human granulocytes (PMN), dissolved these in sample buffer, ran the samples next to each other on an SDS-PAGE gels, blotted the gel to a membrane and probed that membrane with antibodies against the NADPH-oxidase components p67 and p47. Finally, I probed the blot with fluorescently labelled antibodies and scanned it on a Li-Cor Odyssey infrared scanner to detect the signal. As you can see, the neutrophils (PMN) express high levels of both p67 (in red) and p47 (in green) whereas the control PLB cells express nothing and the treated PLB cells express levels comparable to the neutrophils, demonstrating that the experiment was successful. I also probed the blot for β-actin to demonstrate equal loading of the samples. This is a perfectly straightforward result: the protein is either there or not there at all. No discussion about it! However, most blots won’t be so straightforward.

Figure 1. Western blot probed for p67phox (red) and p47phox (green) or actin (lower panel). Loading control on the far left lane with molecular weight bands as indicated in kDa. A sample of human neutrophils (PMN) is shown as a control as compared to non-differentiated PLB cells and PLB cells differentiated with 0.5% DMF.

In most cases, your protein of interest won’t be either there or not there. The level of expression will be reduced or increased by a certain margin. Western blots can be very deceptive in demonstrating such subtle effects, as I will outline below.

Controls

First and foremost in scientific practice come the controls. Your control and experiment samples should have received the exact same treatment to be valid and run side by side on a gel for proper comparison. I know it happens far too often that you go through the whole procedure only to discover a nasty spot in either your sample or control lane. Then you go and repeat the whole procedure, only to have the other lane smudged this time. If only you cropped both pictures and spliced them next to each other, you’d have a perfect result… Unfortunately, this is not allowed. Such practice reeks of scientific fraud and makes it impossible to judge whether or not your result was real, even if you mean well. Nope, my friend, either it’s back to the lab, or show the smudges! There’s nothing wrong with showing smudges, one should not be afraid of that. Smudges are much preferred over fraud!

Detection limits

The second issue to consider before stating whether or not a certain protein has disappeared or not (for example after siRNA treatment) is the detection limit of your protein of interest. The problem with all commercial antibodies is that nobody really knows just how strongly they interact with their target protein. Anybody who’s tried several different antibodies against the same target knows that some will give you a very strong signal and low background, while others will give you a weak signal and high background. Anything in between, in any possible combination, can also be encountered. It’s completely random! This indicates that the strength of the signal on your blot is not so much an indication of relative protein abundance as of antibody quality. Though, of course, protein abundance also helps.

Detection limit becomes especially relevant when your signal is very weak. This means you’re drawing conclusions close to the detection limit of your assay. Thus, when comparing two lanes on a blot, it may seem that in one lane the protein is there and in the other it’s absent, while in reality the difference is no more than 10% or so. What makes the blot deceptive is the detection limit of your protein. It could just happen that you loaded 10% less of your experimental sample and that this is enough to make your protein of interest undetectable, even though it’s still there. If you now compare the control and your experimental lane, it will look like your protein of interest has disappeared completely after treatment and since we usually use highly abundant proteins, such as actin, as our loading controls, this 10% difference in loading will go unnoticed. I see these kind of results quite often with siRNA controls; very weak signals that appear to show a black and white difference. Results collected on old fashioned X-ray film are particularly sensitive to this form of deception, since a pixel on the film is either black or not. The grey values on a film are entirely derived from pixel density. In contrast, results collected with a fluorescence imager are much less sensitive to deception, since every pixel can have a wide range of values. On the Li-Cor Odyssey, for example, a pixel can have a value between 1 and 40,000, providing a huge dynamic range.

Post-processing errors

Every digital image can be manipulated to show only what one desires others to see. Of course, manipulating only part of the image (say the control lane) and not the rest is fraud, but there’s a huge grey area of manipulation that is allowed, but not quite correct. I provide an example in Figure 2. For this experiment, I tried to knock down a gene with siRNA. The blot shows my control sample (Ctrl) and two different siRNA’s (1 and 2). Panel A is what the blot actually looks like after I acquired the data with the Odyssey infrared imager, panel B is manipulated to show what I want to see (protein levels decreased after siRNA treatment) and panel C is my loading control (HSP90). As you can see, the effect of siRNA treatment appears to be much greater in panel B than in panel A, even though both show the exact same blot. What I did to generate panel B was to adjust the image display curves, rather than merely the contrast. You should never, ever do that! Ever. Because by adjusting the curves you’re discarding data you don’t like. You’re telling the program that you don’t care about values above or below a certain threshold, thus you get rid of pixels with very low or very high values. Of course, this eliminates your background, but you also lose information that might be valuable, such as the weak bands visible in panel A below and above the main bands. In addition, you enhance and multiply a small difference to make it seem much greater. This might indeed help you get your work published in Nature, wherein most blots are suspiciously squeaky clean, but it really isn't the way to go.

Figure 2. Control sample (Ctrl) and samples treated with siRNA 1 or 2. A) unprocessed image, B) processed image, C) loading control (HSP90). 

Loading controls

Another obvious problem is with the loading control. As I mentioned before, we tend to choose a highly abundant protein as our loading control, such as Actin. However, if your control is much more abundant than your protein of interest, the result may be highly deceptive. I provide an example in Figure 3. For this experiment, I simply took a cell lysate and diluted this with sample buffer in 10%-steps (thus, lane 1 is 100%, lane 2 90%, 80% etc.). Then I probed the blot with an anti-actin antibody, followed by a fluorescently-labelled antibody and I scanned the blot with the Odyssey Infrared scanner. The nice thing about this technology is that I can actually quantify my signal and draw a graph, as shown in the figure. Now, you’ll notice that the difference between the first 5 lanes is very hard to spot with the naked eye, even though there’s an almost 2-fold difference in loading (100% vs 60%)! In addition, the computer can barely tell the difference between the first 3 lanes, even though I loaded only 80% of the sample in lane 3. Thus, also the intensity of your loading control can be very deceptive and you should be aware that the same phenomenon occurs for every protein you blot for. If you’re trying to draw conclusions outside the linear range of detection, whether by eye or computer, you’re going to be deceived.   

Figure 3. Decreasing amounts of sample were loaded on gel with steps of 10%. The blot was probed for human Actin and data acquired on the Li Cor Odyssey.

Finally, the molecular weight of your controls and experimental samples matter. Not every protein in your sample is going to transfer equally well and molecular weight is an important determinant for transfer efficiency. Large proteins tend to transfer slower whereas small proteins can actually be transferred straight through your blot if your transfer time is too long. Thus, ideally, your loading control and protein of interest should be of both similar size and similar abundance.

Solutions

It’s up to you to provide the right controls for your experiments and to make sure your samples and controls are on the same blot. You can not, and should not, compare samples from different blots as they may have been differently exposed and processed.

You should know the detection limit of your protein of interest and the affinity of your antibody. When you first start using a new antibody, run a control blot with a dilution range of your sample to make sure you’re measuring in the linear range of your protein.

Image manipulation is allowed, of course, you’re already doing it when you’re exposing your film for different lengths of time. However, only manipulate the whole blot and never discard data. Better to show some noise and background. Find some useful hints and tips on this website. Nowadays, I much prefer using fluorescence scanners, such as the Odyssey, the acquisition Western blot data because the scanned imaged can be quantified and I can easily make different exposures of my blot, simply by setting the exposure level. When using the Odyssey, always set the acquisition for the maximal possible exposure, without over-exposing the image (shown as white pixels in the colour view or blue pixels in the grey scale view). That way, you collect as much data as possible and you can always post-process your image to get a prettier picture. More important than a pretty picture, however, is the ability to graph and calculate your data.  

I hope these examples and guidelines give you some idea of how to interpret Western blot results, be it your own data or published work. 

How to make the mutation you want and succeed every time

Good research is highly reliant on good tools. Among the most important tools in molecular biology and biochemistry are mutant versions of the genes we study. These mutants help us understand how the proteins encoded by the genes function, both in vivo and in vitro. Today, I shall discuss the site-directed mutagenesis protocol I've been using to get my mutants. My protocol works. Every. Single. Time.

When it comes to site-directed mutagenesis many people prefer the easy way out and use a kit. The QuickChange kit from Stratagene, for example, is a popular choice. However, this kit is rather expensive, unnecessarily complex and really not as good as advertised. The whole procedure can be performed with materials you probably already have in your lab at a fraction of the cost, if you follow the exact steps outlined below.

1. Primer Design

The first step towards generating a successful mutation is designing your primers. The same rules of thumb apply when designing a mutagenesis primer as for designing a cloning primer: you're looking for a primer that extents at least 9 bases on both sites of the mutation, with a GC content of 40-60% and a melting temperature of about 52-55C. I always use complementary primers, a pair of primers that are each others exact opposites. For example, in figure 1, I show you the primer I designed to mutate an arginine residue, R54, to a cysteine in human Ubiquitin B. I did this by changing a single 'C' in the primer to a 'T', as indicated by the yellow bar. The codon 'CGC' encodes an arginine, whereas ' TGC' encodes a cysteine. Easy as that!

Figure 1. A primer designed  to mutate arginine 54 to a cysteine (R54C) in human Ubiquitin B.

Notice how the primer extents for 10 bases to both side of the mutation? Shorter primers also work, but try to extent it at least 9 bases on both sides of the mutation while keeping the GC content within 40-60% (in this case, it's 52.4%) and the melting temperature between 52-55C (53.5C, in this case). Bear in mind that the mutation you make will not count towards the melting temperature of the primer, as the mutation represents a mismatch! Of course, you can introduce more than one mutation, but the primer will have to be proportionally longer. I've successfully introduced up to seven point mutations with a single primer pair in one reaction using this method.

2. PCR

Once your primer order has come in and you have your template ready, it's time to run a PCR reaction. For site-directed mutagenesis reactions, it's wise to use a reliable polymerase with proof-reading. Several options are available, but my favorite enzyme is Phusion from New England Biolabs. This enzyme is twice as fast as a regular DNA polymerase and a whopping four times as fast as other proof-reading enzymes, such as Pfu. It has, in my experience, also proven to be highly reliable. Using the Phusion enzyme and buffer, set up the following reaction:

200 ng DNA template

125 ng forward primer

125 ng reverse primer 

2 µL dNTP (2.5 mM each)

10 µL 5X Phusion buffer

1 µL Phusion

 x µL water (to 50 µL total)

The ideal amount of template to be used depends on a lot of factors, but I find 50-200 ng to be sufficient. If you're worried about background you can use a little less, but more generally works better.

Set up one reaction with and one without polymerase as a control! This is important, so make sure not to forget this control.

The following PCR program can be used for Phusion (Figure 2), adjust to primer melting temperature (2 degrees below the melting temperature of your primers is ideal) and plasmid size (30 seconds/kb, remember you're amplifying the entire plasmid plus your insert!). 18 cycles is generally sufficient.

Figure 2. General PCR program used for site-directed mutagenesis with Phusion. 

It's best to run the PCR in the morning. The program only takes about 2 hours in total, when using Phusion; enough time for a hearty lunch and a cup of coffee!

3. Analyze Product on Gel 

After the PCR is done, take 10 µL of your product and analyze it on a 1% agarose gel. This is a very important step in the mutagenesis process! If the mutagenesis reaction has worked you'll see a signifcant increase in the amount of DNA in your reaction with polymerase, as compared to your control reaction without polymerase. If you don't see this increase, the reaction hasn't worked and there's no point in proceeding! Have a good look at your primer design or try again with more template DNA. If the reaction has worked, you should see something similar to Figure 3.

Figure 3. PCR product of the mutagenesis reaction analyzed on gel. Samples 1-3 were run without polymerase, samples 4-6 were run with Phusion polymerase added to the reaction.

4. Digest Product with DpnI

If, and only if, you got significant amplification of your template after PCR, you can digest your product with the restriction enzyme DpnI. This enzyme only digests methylated DNA, so your template (which was amplified in and purified from E. coli) gets digested, whereas your mutation-bearing product, formed in the PCR reaction, remains intact. Simply add 1µL DpnI (I get mine from Promega these days, but any DpnI is fine) directly to your PCR tubes and put them at 37C for 2-3 hours. Don't forget them and leave them overnight, as there will be nothing left (I know from experience...).

5. Transform  into DH5a and Mini-prep

After digestion, simply take 5-10 µL of your product and transform into competent DH5a E. coli to amplify the DNA. Any strain of competent cells will work, really. Plate the cells in LB agar plates after the transformation and let them grow overnight at 37C. The plate of the no polymerase control should have no colonies, whereas the other plate should have plenty.

I know from experience that if you digest a product that did not look like it was amplified on gel you will still get no colonies on the no polymerase control plate and some on the other plate. However, none of these colonies will harbor the mutation. I don't know why this is, yet it happens.

The following day, pick some 12 colonies from your plate and grow mini-prep cultures overnight. You'll need enough DNA for sequencing (50 ng/µL) and I usually grow and prep 2 mL culture for high copy plasmids and 4 mL culture for low copy plasmids. I like to use 2xTY medium for my cultures, rather than LB, as the yield is higher.

For your mini-preps you can use any kit you like, but be aware that the popular Qiagen kits are expensive but absolutely no better than any other kit out there. I generally like the kits from Machery-Nagel or, if you really want to safe some money, you can buy the Econo-Prep columns from Epoch Life Sciences and make your own buffers. Mini-prep columns can be recycled and re-used several times, so there's really no use in spending a lot of money here.

6. Sequencing

After you've done your mini-preps, it's time to send out your DNA for sequencing. Most labs these days use commercial sequencing services, and prices have dropped significantly in recent years, so I don't find it necessary to safe money in this step at the risk of losing time. In general, the mutagenesis reaction is rather efficient, but I like to sequence about twelve clones of each reaction to be on the safe side. I find that at least one in four clones generally contains the desired mutation. 

If you don't have access to fast and cheap sequencing facilities, you can also consider designing your mutation so that you introduce a restriction site. That way, you can analyze your clones by restriction digest. However, nowadays, this is hardly cheaper, faster or more reliable than sequencing and you'll have to sequence anyway in the end.

Conclusion

Well, that's it! With this protocol, my mutagenesis reactions always work, provided I designed my primers right and used enough template. If you're introducing multiple mutations with very long primers, you might consider adding 1% DMSO to your PCR reaction. This prevents the formation of DNA super structures during amplification, but also decreases the efficiency of the polymerase. Good luck making some mutants of your own!

Apoptosis vs. Necroptosis

Cell death is a highly regulated process. Ask any cancer or stroke patient. In the former case, too little cell death is causing problems, in the latter it's too much cell death that's doing the damage. Every day, approximately 100 billion (!) cells die in your body and every day, all those cells are replaced. Every day you die a little and you never even noticed.

Most of those cells die by a process called apoptosis; a programmed form of cell death. Every cell is genetically programmed to undergo apoptosis, a sequence of orchestrated events leading to the cells demise, when the cell has either suffered irreparable internal damage or receives an extrinsic stimulus from its environment. The extrinsic signal to die is given when a cell is, for example, infected with a virus, has become old or redundant or has become dislodged from its usual place in the body. The cell is basically told to quietly commit suicide, dismantle itself and allow its remains to be recycled. Cells are equipped with 'death receptors', members of the TNFa super family, that receive the death signal.

As mentioned above, apoptosis can also be triggered intrinsically. When, for example, a cell's genome or essential organelles have suffered irreparable damage a sequence of events leads to the release of toxic proteins from the mitochondria, such as cytochrome c and SMAC, that induce the cell to undergo apoptosis and remove itself from the population.

Extrinsic versus intrinsic apoptosis (image by author)

Apoptosis is a quiet, dignified form of cell death that does not trigger an inflammatory response. Apoptosis depends on the sequential activation of the caspases; a family of cysteine proteases. At the top of the chain are the initiator caspases (caspase-8 and -10 for death-receptor-induced apoptosis, caspase-9 for intrinsically-triggered apoptosis), at the bottom are the executioner caspases (caspase-3, -6 and -7) that dismantle the cell. Diametrically opposed to apoptosis is necrosis; a messy form of cell death -wherein the cell's contents are spilled into the environment- that does elicit an inflammatory response.

Surprisingly, necrosis can also follow a genetically-encoded program, similar to apoptosis. However, programmed necrosis, now widely known as 'necroptosis', does not depend in caspase activity, but on the activity of a kinase: Receptor Interacting Protein Kinase 1 (RIPK1). RIPK1 functions as the initiator of the pathway, while several downstream kinases (most notable RIPK3) serve as the executioners. We don't know much about necroptosis yet, but new findings on this form of cell death are published almost daily. What we do know for sure is that caspase activity is essential to prevent it. Necroptosis only occurs in the absence of caspase activity (for a free review by some friends of mine, see here).

The initiators of death-receptor-dependent apoptosis, caspase-8 and caspase-10, have both been shown to cleave and inactivate RIPK1 (reviewed by me here). Thus, if those caspases are activated, RIPK1 is inactivated. The opposite is also true: When the gene for caspase-8 is knocked out in mice (mice don't have the gene for caspase-10), the embryo deficient in caspase-8 dies on the eleventh day after gestation (Varfolomeev et al. 1998). This is a crucial day in the development of the murine embryo, since at that time the embryo's own blood circulation kicks in. In the absence of caspase-8 activity, the hearth and blood vessels of the embryonic mice fail to develop. Experiments with conditional knockout mice, mice that are only deficient for caspase-8 in certain tissues, have revealed that caspase-8 activity is also essential for the development of the immune system (see Kang et al.).

This failure of caspase-8 deficient embryos to develop, is entirely due to the activity of RIPK1 and its downstream effector RIPK3. Knock either one of these genes out concomitantly with caspase-8 or the adapter protein FADD (essential for the activation of caspase-8) and the mouse develops just fine (here, here, here and here; only the last one, the one I'm on, is free) . Or at least; it develops past this crucial stage, past day 11, for knockout of RIPK1 is lethal in itself. RIPK1 has pro-life as well as pro-death functions, but knockout of RIPK3 is relatively safe. Mice deficient in RIPK3 as well as caspase-8 develop into relatively healthy individuals. They still have some problems, of course, caspase-8 and RIPK3 are not entirely useless genes you can just dispose of. They have problems dealing with viral infections, for example, and their T cells proliferate unchecked. They may have other problems too, that haven't surfaced yet.

Thus, caspase-8 has a crucial pro-survival role in shutting off RIPK1 and preventing it from inducing necroptosis. But how, then, does a cell wherein caspase-8 is activated not die by apoptosis instead? How does it live to develop into a healthy mouse or human? Caspase-8 activates through dimerization; two molecules of caspase-8 are forcefully brought together to form an active complex. The previously mentioned adapter protein FADD is essential for initiating this process of dimerization, but recent evidence has shown that once a few dimers are formed around clusters of FADD, more caspase-8 dimers can form independent of FADD. An important clue comes from the observation that caspase-8 does not only activate when it dimerises with itself to form a homodimer, but can also when it forms a dimer with its cousin, FLIP (FLICE-like Inhibitory Protein), to form a heterodimer. FLIP is similar to caspase-8 but has no protease activity, it is an inactive caspase homologue. The heterodimer is active, but has a restricetd substrate repertoire; it cleaves the pro-apoptotic substrates of caspase-8 with very low efficiency, while it cleaves the non-apoptotic substrates of caspase-8 just as efficiently as the homodimer. I recently published a very readable review on the proliferative versus the apoptotic functions of caspase-8, find it here.

Apoptosis vs. Necroptosis vs. Survival  (image by author)

Now, this is not the end of it. There is much more to RIPK1 signaling than necroptosis; it plays an important role in immune activation and development too. In addition, recent evidence suggests that direct cleavage of RIPK1 cleavage by caspase-8 may not even be the key to prevention of necroptosis. Instead, caspase-8 may cleave CYLD, a de-ubiquitinating enzyme and an important regulator of RIPK1 activity. As long as CYLD is active, RIPK1 can promote necroptosis but if CYLD is inactivated RIPK1 is more likely to promote cell survival. However, caspase-8 is very bad at cleaving either CYLD or RIPK1. The paracaspase MALT1 can also cleave CYLD, an event that is crucial for the activation of T cells, but it is not so very good at it either. Could there be another caspase, downstream of caspase-8, that cleaves and inactivates RIPK1? Does cleavage of RIPK1 really lead to its inactivation or do the two fragments gain a different function? Does the caspase-8/FLIP heterodimer have other substrates, besides RIPK1 and CYLD? These and other important questions are currently under investigation. We're not even sure yet what the relevance of necroptosis is for either normal human physiology or pathology.

Surely these are exciting times for the fields of cell death and inflammation! I will use this blog to review the latest findings in these fields, both by myself and by others. I hope to attract opinionated readers both inside and outside the fields and get some discussions going.