Author: Aaron

When to use nitrocellulose vs PVDF membranes

Categories
Transfers Western Blotting

While we might not like to admit it, us scientists are creatures of habit, reducing variables to ensure our hypothesis testing is as accurate as possible rapidly creeps into our day to day lab tasks. In some ways this can be good, with standard operating procedures and recipes being handed down through the lab to ensure consistency, but in other ways it can be limiting. Today I’m going to talk about one area that’s particularly close to my heart… Western blotting membrane choice.

Female scientist using tweezers to look at membrane
Using pre-cut membranes saves you time in the lab.

Now I’ll freely admit it’s not the coolest of topics, but it’s something worth considering, especially if you’re struggling with your Western blotting. I went through a period of using exclusively polyvinylidene difluoride (PVDF) membranes as that was what I’d always used. One protein though was giving me a lot of trouble, lots of background regardless of antibody used, block or transfer conditions. At the end of my tether and tearing my hair out I borrowed some nitrocellulose membrane as a last resort, and it worked straight away. So sometimes being flexible can be beneficial as well.

Table of contents

How to choose the right membrane for Western blotting

In the majority of situations, either PVDF or nitrocellulose membranes will probably work well for you, but there are certain circumstances when one is preferred over the other. Choosing between membranes depends on your detection method, whether you intend to strip and reprobe, and your target protein(s) (Table 1).

While you can use both nitrocellulose and PVDF membranes for your Western blot analysis, nitrocellulose membranes are ideal for detecting low molecular weight proteins and PVDF membranes are more suitable for detecting higher molecular weight proteins. Both PVDF and nitrocellulose membranes are compatible with chemiluminescence-based Western blotting. For fluorescence-based Western blot detection, we do not recommend using a nitrocellulose membrane, due to its high autofluorescence.

Two most common types of membranes used in Western blotting

Sometimes your experiment may require a more sophisticated membrane than standard. Using the optimal membrane for your Western Blot application can be critical to your experiment’s success.

Table 1. How to choose between nitrocellulose and PVDF membranes for Western blotting

NitrocellulosePVDF
Repeated re-probesAppropriatePossible, but sensitivity is lost
Small proteins<25 kDa>25 kDa
DetectionGreat to use for chemiluminescent and fluorescent detectionUse dedicated low-fluorescence PVDF membranes for chemiluminescence detection
Protein bindingHydrophobic proteinsHydrophobic and dipole proteins
Sample concentrationLow abundance
80-100 µg of protein/cm2		Higher abundance
150-200 µg of protein/cm2
AutofluorescenceLowHigh with standard PVDF membranes

Nitrocellulose Membranes

Nitrocellulose was the first Western blot membrane and is still widely used today. Ready to use and easily hydrated nitrocellulose membranes rapidly and readily bind with proteins out of the box and are often more affordable than other membrane types. However, they often lack durability making stripping and re-probing more difficult.

Two packages of nitrocellulose membranes
Azure offers multiple sizes of nitrocellulose membranes to accommodate larger and smaller proteins. Each package comes with ten membranes in two pore sizes: 0.45 μm and 0.22 μm.

Why are nitrocellulose membranes used in Western blotting?

Nitrocellulose membranes are extensively utilized in Western blotting due to their superior protein-binding affinity, compatibility with multiple detection methods (chemiluminescence and fluorescence, to name a few), and efficacy in immobilizing proteins, glycoproteins, or nucleic acids.

Polyvinylidene difluoride (PVDF) Membranes

PVDF membranes are more robust, but a bit more complex to work with. They need to be activated in methanol before they can efficiently bind proteins, and often also require methanol and SDS in the transfer buffer.

Package of pre-cut PVDF membranes for Western blotting applications
Pre-cut PVDF membranes can be used for a variety of Western blotting applications to save you time in the lab. Consider using theselow-fluorescence PVDF membranes to reduce background noise and improve sensitivity.
Purchase Membranes

Why are PVDF membranes used in Western blotting?

PVDF membranes are utilized because they are robust and can bind more protein than their nitrocellulose counterpart. They are ideal for low abundance proteins. PVDF membranes offer increased sensitivity, with the tradeoff of being more prone to showing non-specific background staining.

Other types of membranes

While most labs use PVDF or nitrocellulose membranes, other options are available. These include nylon or cellulose membranes. However, these are usually used only in specific circumstances. I hope after reading these tips we were able to help you pick the right membrane for your assay. It’s always worth considering a change if you’re not getting the bands you expect. Make sure you have everything you need for your next Western blot, by checking out this Western blot resources list.

If you’re still having issues selecting the right membrane, fill out the form on this page and one of our scientists will help you narrow it down to the right membrane.

FREE WESTERN BLOT eBOOK

New to Western blotting? Need to troubleshoot your Western blot?​ Want to brush up on Western blotting best practices? Claim your free Western Blotting eBook!
Download eBook

More posts on Western blotting:

Shop Membranes and Reagents for Western Blotting

Multiplex Western Blots: 6 Strategies that Work

Categories
Multiplex Western Blotting

Are you having problems with your multiplex fluorescent Western blots? In this post, we’ll cover the exact protocols in more detail in some of our application notes that cover phosphorylation/total protein detection, and single step normalization.

Detecting one protein, two proteins, three proteins? How about FOUR proteins on the same Western Blot? Multiplexing your Western blots allows you to save money on each experiment.

Here at Azure Biosystems we’re great believers in the power of fluorescent multiplexing in Western blots, which has driven a lot of the development of our line of Azure imagers. But we recognize that taking the step from a standard HRP/chemiluminescent-based approach can be daunting. Keep reading for 6 tips to make the switch as easy as possible.

Azure fluorescence capabilities vs. competitor fluorescence, 3 proteins, plus total protein staining, and multiplex fluorescent Western blot

Browse our 6 multiplex Western blot expert tips

AZURE EXPERT TIP #1: Increase your concentrations

The concentration of both the primary and secondary antibody required may be increased compared to chemiluminescent Westerns depending which imager you’re using. In more modern or laser-based imagers, this effect may be less marked.

AZURE EXPERT TIP #2: Test individually first

Going hand in hand with our first tip, it always makes sense to test your antibodies individually first. That way you can determine optimal concentrations, factors contributing to background or lack of specificity in a simpler environment, rather than try to unpick 3 different primary and secondary antibodies at once.

AZURE EXPERT TIP #3: Use adsorbed secondary antibodies

Although it sounds simple, many people don’t consider that they are now adding multiple antibodies from multiple species onto a single blot. If you work with multiple fluorescence in other fields you’ll already be aware of the importance of cross-adsorbing secondary antibodies to reduce inter-species cross reactivity. But many people don’t consider this for Westerns, it’s worth checking out your antibodies to ensure they meet the grade.

Multiplex fluorescent Western blot
Multiplex fluorescent Western blot imaged with an Azure Imager using Cy3 and Cy5.

AZURE EXPERT TIP #4: Expand the spectrum with multiplexing

Three-color Western blots are exciting, but what about five colors? With near-infrared capability, the number of spectrally distinct peaks that can be isolated can be increased greatly. Obviously, this may require a bit of work up to get the antibodies optimized, but imagine the time and cost savings generated by performing one Western for five proteins.

The Sapphire Biomolecular Imager is an imager capable of detecting up to four proteins on the same Western blot, which allows you to save money on reagents each time you do an experiment. It uses 4 fluorescence channels to easily quantify overlapping bands. Read more on multiplex protein detection here. 

Since the release of this blog post, the Azure Sapphire has been succeeded by the new Azure Sapphire FL, which was designed to be the flexible choice in bringing precise quantitation of nucleic acids and proteins.

AZURE EXPERT TIP #5: Check your membrane

Some membranes will auto-fluoresce when exposed to UV light generating a high background signal, although more and more fluorescent safe membranes are being developed. We would also recommend switching to using PVDF membranes from nitrocellulose due to its increased sensitivity, as we discussed previously.

Package of pre-cut PVDF membranes for Western blotting applications
PVDF membranes reduce background noise for improved sensitivity.

AZURE EXPERT TIP #6: Choose the right channel for your protein

Unfortunately all detection channels were not created equally. For standard fluorescence use blue to detect your highest abundance protein, green the middle and red for your lowest abundance protein. If introducing NIR the excellent sensitivity and low background achieved with these fluorophores also makes them ideal for low abundance proteins.

Want to read more? Check out this paper from the National Institute of Health that utilizes multiplex Western blots using microchip electrophoresis.

Have more questions or want to learn more about multiplexing and how it can improve the way you research? Fill out the form on the left- we’ll be in touch.

FREE WESTERN BLOT eBOOK

New to Western blotting? Need to troubleshoot your Western blot?​ Want to brush up on Western blotting best practices? Claim your free Western Blotting eBook!
Download eBook

Which protein assay is best for you?

Categories
Protein Assays Quantification

Have you ever spent ages correctly quantifying your protein samples, casting the perfect gel, efficiently transferred, probed and eventually come to the visualization stage only to be presented with a loading control like this:

Factors to consider when choosing the right protein assay

Before you throw your blot out with the bathwater and repeat the experiment, you should consider how your choice of protein assay can interact with your samples. We’re here to help you detect the presence of protein.

The two most common methods for the colorimetric detection and quantitation of total protein are

  1. Protein-copper ion-based and
  2. Protein-dye binding assays.

We’ll cover both types and more in this blog post.

Major methods of determining protein concentrations

Most modern protein assays are based on two colorimetric assays. The first relies on the interaction of proteins with copper and the subsequent reduction of these copper ions (Biuret and its derivations BCA and Lowry). The second on direct alterations to Coomassie Blue dye due to its interaction with proteins in solution (Bradford). Both the protein sample’s nature and presence of non-protein agents can significantly affect the results of your experiment.

Keep in mind most colorimetric assays require at least 0.5µg proteins in order to produce reliable estimations. You will need proper instrumentation to measure the absorbance. Using an absorbance microplate reader with a highly sensitive and reactive read speed and high accuracy rate will allow you to quickly get your results. We recommend using the Ao Absorbance Microplate Reader, which a read speed of <6 seconds, and an accuracy of >0.005 +/- 1% (0-3) OD. Using this reader, you can comfortably and quickly run any assay using an intuitive 7” touchscreen, without the need for a separate computer.

Table 1. How to choose a protein assay

ConditionChoose this protein assay
You see odd looking results even though you're sure your quantification is accurate and your loading control is correctLook at your protein assay again
Your extraction buffers contain reducing agents such as DTT or BMEBradford protein assay
Your samples contain detergentsEither BCA or Lowry assay
You're working with high concentration samplesBiuret assay. Keep in mind it is often preferable to simply dilute high concentration samples down.

Types of Protein-copper Assays

Protein-copper assays based are divided into two groups:

  1. Assays that detect reduced cuprous ions (Cu+) and
  2. Assays that detect unbound cupric (Cu2+) ions. Bicinchoninic acid (BCA) may be used to detect cuprous ions.

Biuret Test

The biuret test (also called Piotrowski’s test) was first described in 1833, although its usage as a protein assay came much later. Given its relative insensitivity (how many of us are using samples in the range of 5 to 150 mg/mL) and lack of reactivity with single amino acids or small peptides, the biuret assay is ideal for high protein samples, such as tissue.

A test tube containing a clear violet solution
A positive biuret assay sample. Image c/o Philip Evans

At its most simple an aqueous protein sample is treated with as strong base (typically sodium hydroxide NaOH) and a copper(II) sulfate solution. When proteins are present the copper(II) binds with nitrogens present in the protein sample. These liberated copper(II) molecules can now be reduced which in turn changes the color of the solution from a pale blue into a nice shade of purple as seen below. As the amount of reduced copper ions are correlated with the amount of protein present in the sample, and in conjunction with a standard curve, it is then possible to accurately quantify proteins within the range of 5 to 150 mg/mL.

There are two derivations of the biuret assay which be used to dramatically increase assay sensitivity: the BCA and Lowry assays.

Bicinchoninic acid (BCA) Assay

The BCA assay is a high-throughput colorimetric assay that gives accurate quantitation. In the BCA assay the copper ions generated with through interaction with proteins react with BCA, producing a strong violet or blue-ish color. The difference between biuret assays and BCA assays, however, is the formed BCA/copper complex absorbs light much more strongly than those generated by the biuret assay. For this reason alone, the assay’s sensitivity is greatly increased.

With proper sample preparation and accurate standard curves, it is possible to assay samples accurately within the 0.001 to 2 mg/mL protein concentration range.

It’s important to note the color density is affected by the presence of asparginine, tyrosine, cysteine, tryptophan, and histidine. Avoid EDTA (>10 mM) and other strong chelating agents with BCA assays. The color produced (absorbance) can be read at 650nm to 750nm and is proportional to the amount of peptide bonds. BCA assays are incompatible with reducing agents and copper-interacting reagents like ammonia.

Lowry Assay

The Lowry assay is an earlier derivation developed by Oliver Lowry in the 1940’s. In a similar fashion to the BCA assay, the Lowry assay relies on the reaction of copper ions, produced from the interaction of copper(II) sulfate with proteins present in the sample, with a further reagent. In this case Folin–Ciocalteu reagent with the sample becoming blue in color when higher levels of protein are present. This assay can be a tad tricky, as it’ll take practice to add certain components need to be added at specific time points. You will be able to measure the Lowry method between 650 nm and 750 nm without loosing color intensity. 

From left to right, decreasing protein concentrations as assessed by the Lowry protein assay. Image c/o Imgur

The Lowry method is sensitive in the range 5 to 150 μg protein per assay. It is often noted as more consistent than the Bradford assay. Because of its sensitivity to low molecular weight peptides and proteins, the Lowry method is able to more accurately evaluate the increase of purity during fractionation.

It is important to note that the Biuret Test, BCA Assay, and Lowry assay are sensitive to reducing agents often used in sample preparations, such as dithiothreitol (DTT) or beta-mercaptoethanol (BME). These reducing agents can inflate protein concentration values by increasing the reduction of copper ions.

Types of Protein-Dye Assays

Protein-dye assays are not sensitive to reducing agents; however, they are sensitive to detergents, such as Triton, Tween or sodium dodecyl sulfate (SDS).

Bradford Assay

The Bradford assay is fast to perform and gives accurate quantitation. It uses the same amount of protein as the Lowry assay. The Bradford assay relies on an absorption shift by Coomassie Blue when it binds to proteins present in a solution. Under acidic conditions unbound Coomassie will present its typical blue-green color with a maximal absorbance of 465 nm.

So… Which protein assay to chose?
  • If you see odd looking results even though you’re sure your quantification is accurate and your loading control is correct: consider looking at your protein assay again
  • If your extraction buffers contain reducing agents such as DTT or BME: consider the Bradford protein assay
  • If your samples contain detergents: BCA or Lowry assay may be preferred.
  • If you’re working with high concentration samples: the biuret assay may be used. Keep in mind that it is often preferable to simply dilute high concentration samples down.

However, under the same conditions, Coomassie readily binds non-covalently with the carboxyl groups of proteins which induces a conformational shift allowing for a further ionic bond to form between the protein and the Coomassie dye. This binding alters the absorbance profile of the dye and changes the color from brown to blue, which can be measured at 595 nm. By measuring the shift from 465 nm to 595 nm and in conjunction with an appropriate standard curve, accurate protein quantification is possible.

The main limitation of Bradford assays is that it is not compatible with most detergents. Another major drawback of the Bradford protein assay is that it does not work if the sample is basic, or surfactants or detergents are in the sample. 

Still unsure about which protein assay to choose? We’re here for you! Fill out the form on this page to ask a question.

How to Get Consistent Answers to a Rainbow of Assays

Save time with ELISA, Bradford, and Lowry assays with wavelength ranging from 340-750nm. Each microplate reader comes with an 8-position filter wheel with 8 filters allow you to measure the color produced by your assay of choice.
Azure Biosystems Ao Absorbance Microplate Plate Reader
Ao Absorbance Microplate Reader
Get Quote

The evolution of the ELISA and other immunoassays

Categories
Immunoassay In-cell Western Multiplex Protein Assays

If you’re reading this blog there’s a high chance you’re using antibodies in your work, in some form of immunoassay. However, many of us rarely stop and think about where these antibodies come from and how their development has been fundamental for the evolution of life sciences.

A brief history of the antibody

Antibodies as an idea were first hypothesized in the 1890s when Shibasaburō described what we would now call antibody activity against diphtheria and tetanus toxins, suggesting that some molecule in the blood was driving this effect. This led to Paul Ehrlich to propose the theory for antibody-antigen interaction and indeed coin the term antibody, although it was several years before this usage was accepted.

The first molecular evidence of these interactions occurred in the 1920s when Heidelberger and Avery demonstrated that antibodies were indeed proteins produced by cells and that they could be used to isolate their antigen out of a solution. This led to a rapid expansion of the field with the seminal work by Linus Pauling in the 1940s confirming the lock-and-key theory and showing the antibody-antigen interactions are more dependent on their shape rather than their specific chemistry. At the same time Astrid Fagreaus discovered that it was circulating B cells which were the major source of antibodies, and together these two discoveries kick-started the antibody industry as we know it today.

Antibody chains
Image published under the GNU Free Documentation License, Ver 1.2

Building on from this significant work has been performed in identifying and refining our understanding of antibody structure with the discovery of the heavy and light chain structure and the immunoglobulin groups A, D, E, G and M. Which are driving the development of multiplex assays further improving our technical capabilities.

So now that we have an understanding of the history of antibodies, how do they relate to immunoassays including perhaps the most currently widely used enzyme-linked immunoassay (ELISA)? Enzyme immunoassay (EIA), commonly referred to as enzyme linked immunosorbent assay (ELISA), is a vital diagnostic tool used to identify specific antibodies or antigens in a sample. While there are many variations of immunoassays (such as ELISAs, radio-immunoassays, blood typing assays etc.), they all rely on two specific antibody functions, capture and detection.

Capture or Detection

Immunoassays rely on the sensitive and specific detection of antigens by antibodies; however, such assays also need to be repeatable and relatively quick to perform. These two contrasting aims and the varying affinities of antibodies for their antigens allows us to utilize antibodies with different capabilities for different immunoassays, or steps within an an immunoassay.

Antibodies with a very high affinity and selectivity for their antigen are ideal for “pulling” and “holding” their antigen out of a mixed solution, allowing non-specific proteins or other molecules to be washed away. Antibodies like this make ideal capture antibodies and are usually fixed onto a surface allowing sample to be added and washed away, as in an ELISA. Such antibodies are usually monoclonal, providing a high degree of specificity with the potential to under detect some samples.

Conversely, antibodies with a quicker binding rate, but with a reduced affinity, make ideal detection antibodies working on samples which have already been immobilized. As they don’t require high affinity, these antibodies are often polyclonal, which allows for a greater detection range often at a cheaper price. However, this is often dependent on the way in which the sample has been immobilized.

These two families of antibodies can then be applied in a variety of ways depending on assay requirements.

Direct vs. Indirect vs. Sandwich Detection Methods

The ELISA technique has multiple configurations, including: direct ELISA, indirect ELISA, sandwich ELISA, and competitive ELISA. Each variation has distinct benefits depending on the intended application and implementation scenario.

Table 1. Immunoassay types

ELISA typeHow it works
Direct detectionSubstrate is immobilized onto a surface and then primary detection antibodies are added. These antibodies are directly conjugated with a measurable signal, such as a fluorescent or radioactive probe, or an enzyme such as HRP which can drive a colorimetric change.
Indirect detectionAdds a conjugated secondary antibody to the direct detection method. Secondary antibodies are usually designed to target the constant region of the primary antibody, but does so over a variety of sites meaning that a single primary detection antibody can be bound by several conjugated secondary antibodies.
SandwichThe capture antibody is immobilized onto a specific surface and exposed to the sample. This can be washed off to remove all unbound and any weakly bound product. The sample is incubated with a detection antibody, which can display much reduced specificity but increased binding efficiency to amplify signal.
  • Direct detection immunoassays (Figure 1) are the most rapid but are less stringent and so are ideal for tests which require a rapid readout, but where ultimate accuracy is not required. In these assays the substrate is immobilized onto a surface and then primary detection antibodies are added. These antibodies are directly conjugated with a measurable signal such as a fluorescent or radioactive probe, or an enzyme such as HRP which can drive a colorimetric change.
  • Indirect detection (Figure 1) immunoassays improve upon the specificity of direct detection immunoassays by introducing a conjugated secondary antibody. This step also has the added benefit of allowing for signal amplification, which is especially important for low abundance samples.
Difference between Direct vs Indirect ELISA
Figure 1. Direct detection [left] immunoassays are the most rapid but are less stringent and so are ideal for tests which require a rapid readout, but where ultimate accuracy is not required. Indirect detection [right] immunoassays improve upon the specificity of direct detection immunoassays by introducing a conjugated secondary antibody.
  • Sandwich immunossays provide the greatest level of sensitivity and detection. The capture antibody is immobilized onto a specific surface and exposed to the sample. This can be washed off to remove all unbound and any weakly bound product. The sample is incubated with a detection antibody, which can display much reduced specificity but increased binding efficiency to amplify signal. This detection antibody can itself be conjugated or as a further step a secondary antibody can be used.

The downside to the sandwich method is that with all the required steps, the assay length is increased. The difficultly lies in the finding antibodies that bind to the same target efficiently (and without interference) of each other.

The vast majority of immunoassays on the market today use these basic techniques. Understanding how they interact is key for the development of future assays, including the exciting development of in-cell Western blotting, sometimes called cell based ELISA or in-cell Western, which I hope to cover in a later post.

Loading controls and antibody validation for Western blot quantification

Categories
Controls Quantification Western Blotting

Today’s post is inspired by a conversation I had with a colleague when teaching them SDS-PAGE and Western blotting. Being well versed in qRT-PCR (R) and flow cytometry (R), they were particularly interested in how and why certain loading controls were used when Western blotting, and what other techniques were used to validate loading, changes in protein expression and antibody specificity. After explaining to them, I thought I would share with you all too.

Preparing samples

Loading controls ideally form part of a multi-step process to ensure a standard amount of protein is loaded onto a gel for each sample analysed. The first step upon isolating samples is often to perform a protein assay. The BCA and Bradford assays are two of the most popular assays because they allow for standardization of sample concentrations before beginning the assay. While a great first step, there are several issues with the technique: the main one being assay interference from contaminating substances or those used to treat, prepare or isolate samples. For this reason, other pre-separation standardization techniques should be considered alongside proteins assays, such as working with a constant sample size or cell number.

Once the sample has been separated and transferred an often-overlooked, control is to perform a membrane stain, which I discussed in a previous post. Ponceau S staining is a great validation technique to check transfer efficiency, but samples should also look broadly similar if equal amounts of sample have been loaded. With today’s modern image acquisition hardware and analysis techniques, it is also possible to perform total-protein (R) based quantification using Ponceau S (R), Coomassie blue (R) and stain-free methods (R).

  • Azure Quick Tip #1: Ponceau S staining is performed after transfer

    It works with both PVDF and nitrocellulose membranes and is also easily washed off it is hard not to recommend it above other techniques.

e-cad-actin-88

Loading controls

So, onto loading controls. The main purpose of loading controls in Western blotting is to ensure the lanes in your gel contain the same amount of sample and that when you transferred onto the membrane, it was done equally. To confirm that alterations in protein levels are due to effective “knock-down” or “knock-out” and not general changes in protein expression, loading controls are essential.

Most of us have a favorited which we use rain or shine with beta-actin and GAPDH being perennial choices. However, consideration of tissue type, protein of interest localization and size should always be performed, as there are numerous examples in literature that demonstrate how loading controls can and do vary widely (R,R,R).

Here is an excellent resource which details some example loading controls along with their molecular weight, and cellular localization. Keep in mind it is also always worth reading the recent literature to assess previous studies.

  • Azure Quick Tip #2: By performing all three control steps during sample preparation and analysis Western blot quantification should become a breeze.

    This is especially true if you're using a bioimaging system. These systems open the door to multiplex labelling and allow loading controls and the sample of interest to be visualized simultaneously, and total-protein quantification and correction.

Antibody controls in Western blotting

Let’s move onto antibody controls. In Western blotting this area is often overlooked in relation to other antibody based techniques such as FACS or immuno-histochemistry (IHC).

This is partly due to the separation of proteins during electrophoresis which makes determining antibody specificity much easier. It is always worth considering introducing controls for important validations, particularly for Western blots you want to quantify.

Table 1. Four types of Western blot controls include positive, negative, secondary antibody, and isotype controls

Control typeWhat it is and when to use
Positive controlWe use a lysate made from a cell line or tissue sample that has the protein we are trying to detect to test if the antibodies are working properly.
Negative controlTo make sure a test is accurate, we use a negative control. This is a lysate from a cell line or tissue sample that doesn't have the protein we're detecting. Negative controls check for antibody specificity, while also revealing nonspecific binding and false positives.
Secondary antibodyWe only add the second antibody on the membrane to check for antibody specify. In this step, we are able to look for nonspecific binding and false positive results cause by the secondary antibody.
IsotypeIsotype controls help make sure the primary antibody is specifically binding solely to the constant region (Fc) of the primary antibody. Use this for lysates that may be rich in immune cells, or heavily degraded samples.

Positive and Negative Controls

Positive controls are the most widely used in Western blotting and there are several types available. Full length recombinant proteins make an ideal positive control as they should separate identically to the natural protein and can also be used as standards for further quantification if a standard curve is generated.

However, recombinant proteins are not available for proteins and the cost of generation can be prohibitive. In these instances positive cell or tissue samples can be used, including those which have been modified to over-express the protein of interest.

  • Azure Quick Tip #3: Working in the other direction negative control samples can also be used, while a complete blank can be used a better control is a lysate in which the protein of interest is lacking.

    This can be sourced naturally if the protein of interest is relatively tissue specific, or can be generated by knockdown or knockout.

Secondary-only and Isotype Controls

Secondary-only antibody and isotype controls are a mainstay of IHC and FACS; however they are not widely used in Western blotting. Secondary-only antibody controls are performed, as the name suggests, by omitting the primary antibody step from the protocol. This will determine if any non-specific binding or false positives are arising from the secondary binding directly to proteins or protein fragments.

Isotype controls perform a similar function, in checking for non-specific binding, but rather focus on the binding of the constant region (Fc) of the primary antibody, rather than the paratope. This test is particularly important for lysates which may be rich in immune cells which often feature Fc receptors, or heavily degraded samples where non-specific biding can occur.

So to summarize, there are numerous controls that can be performed to ensure your Western blot is optimized for quantification and that you’re observing truly specific binding. While obviously important in its own right, evidence of proper controls is also of growing interest to journals seeking the best quality, repeatable work. So rather than just doing a quick beta-actin strip and re-probe, you should consider some of these additional techniques as well.

FREE WESTERN BLOT eBOOK

New to Western blotting? Need to troubleshoot your Western blot?​ Want to brush up on Western blotting best practices? Claim your free Western Blotting eBook!
Download eBook

Antibody Labeling in the Lab

Categories
Antibodies Multiplex Western Blotting

This blog post will cover antibody labeling in the lab. While imaging technology is advancing rapidly offering us scientists new and interesting ways to image our blot (fluorescent, NIR, multiplexing) the actual technique has remained remarkably similar. The majority of us will block the membrane, probe with a primary followed by a labelled secondary antibody, so called indirect detection. Direct antibody labelling allows the use of antibodies from the same species or of the same isotype and reduces the potential for cross reactivity.

Simple antibody labeling protocol
Simple antibody labeling protocol

With horse radish peroxidase (HRP) conjugated secondary antibodies and chemiluminescent imaging multiplexing was possible but only under certain conditions. Requiring two or more highly specific primary antibodies raised in differing species, or with significantly differing isotypes. With non-murine monoclonal antibodies relatively rare, achieving such specificity was often difficult. If antibody pairings could be identified then species or isotype specific secondary antibodies would then be used; however, this methodology requires a lot of validation to confirm that no cross reactivity is occurring.

Difference between Indirect vs. direct detection in Western blotting
Figure 1.3. Indirect vs. direct detection

With fluorescent-capable imaging systems, such as the Azure Imaging Systems, and the development of a wide range of fluorescent conjugated secondary antibodies, multiplexing suddenly became more feasible as it was possible to identify based on fluorescence where multiple secondary antibodies were binding on a single blot. The development of near-infrared (NIR) imaging systems and antibodies improved this further by expanding multiplex capability while also significantly reducing sample and membrane auto-fluorescence.

However, imaging systems have to some extent advanced beyond the capabilities of traditional indirect detection methods. For example, the Azure 600 can spectrally separate four differing wavelengths (490, 550, 700 and 800 nm). However, identifying four primary and four secondary antibodies which work together in multiplex requires a significant investment in time. In many instances, certain combinations are just not possible to run together.

Example of a four-color Western blot imaging bands in 490, 550, 700 and 800 nm wavelengths, generated using AzureSpectra Secondary Antibodies and AzureSpectra Antibody Labelling Kit Labelled recombinant proteins.

Direct Western blotting

Direct Western blotting is a way to reduce this time investment, and is been driven by developments in fluorescence and NIR antibody design and conjugation. As opposed to indirect detection where a primary antibody binds to the sample, which is in turn bound by a conjugated secondary antibody; direct detection skips the middle-man and uses a conjugated primary antibody and no secondary.

While this technology has long been possible you can imagine why it was of limited use when chemiluminescence reigned supreme, multiplex separation would not be possible and the benefits of indirect detection (secondary antibody mediated amplification increasing sensitivity and specificity). However, using our example above where we had four primary and four secondary antibodies, by switching to a direct system we can significantly reduce the complexity of the system. However, many labs already have well validated antibody panels and it might not be possible to purchase these antibodies pre-conjugated and furthermore conjugates are typically limited to FITC/Cy5 channels.

Perform multiplexing assays quick and easily. Fluorescent Western blot imaged with Azure c600. Secondary antibodies were labeled with the AzureSpectra labeling kit: anti-rabbit with AzureSpectra 700, anti-mouse with AzureSpectra 550, and anti-rat with AzureSpectra 800. Primary antibodies used were rat anti-tubulin, rabbit anti-beta actin, and mouse anti-GAPDH.

Primary antibody conjugation

In order to aid researchers who have a well validated panel of antibodies, but who are interested in exploring multiplexing with directly labelling we, and others, have developed antibody labelling kits such as the AzureSpectra Antibody Labelling Kit. The kits were specially designed to allow researchers to specifically conjugate their validated primaries with a wide variety of fluorescent dyes. AzureSpectra Antibody Labeling kits dramatically ease the transition towards multiplexing and reduce assay time.

Direct antibody labelling allows researchers to have personal control of antibody conjugation provides the utmost flexibility: the short and easy to follow protocols allow researchers to label specific aliquots of antibodies for various techniques including Western blotting, flow cytometry and immunofluorescence.

If multiplexing or antibody labelling sounds interesting to you and you’d like to discuss further, fill out the form on this page and one of our scientists will be in touch!

Shop Secondary Antibodies and Labeling Kits

Trouble-free Transfers

Categories
SDS-PAGE Transfers Troubleshooting Western Blotting

In a previous post I talked about bendy bands, a common problem with improper gel casting and running. However, whilst transferring would seem to be an easier protocol, things can and do go wrong. Below I cover several helpful hints to get you back on track to detecting your proteins after transferring.

No bands after the transfer step: what does it mean?

Unfortunately, the first time you’ll likely consider that your transfer hasn’t worked is when you’re cursing in the darkroom as bands fail to materialize (or in front of your digital imager in the lab, but you’re less likely to be cursing there). That’s probably what led you to this post. 

Here are three steps you can try to eventually detect your proteins.

STEP 1: Simple transfer check

Many of us use a pre-stained ladder in our gels, a simple easy check of transfer efficiency is to check your gel for any residual dye. Your ladder should have transferred across in its entirety, if it hasn’t, something may be wrong in your protocol. If this is the case: try a longer, cooler transfer and remake your buffers.

QUICK TIP: If you see absolutely nothing, check the polarity of your transfer. Run the proteins out of your gel and into the filter paper, rather than the membrane is an easy mistake. But usually one you’ll only make once. Consider color coding the sandwich equipment and leads to ensure the correct polarity at all times.

If you see absolutely nothing, check the polarity of your transfer. Running the proteins out of your gel and into the filter paper, rather than into the membrane is an easy mistake. But usually one you’ll only make once. Consider color coding the sandwich equipment and leads to ensure the correct polarity at all times.

STEP 2: Perform a Detailed Transfer Check using Ponceau Stain

If you want more information about your bands, using Ponceau stain is a great, simple step to validate your transfer. Use Ponceau stain immediately after transferring your membrane. This will be a great way to check for efficient transfer across the length of your lanes.

Comparison of protein transfer using ponceau stain from Azure Biosystems
Ponceau stain was used to check protein transfer before Western blotting to detect solubilization of a protein of interest from inclusion bodies. Duplicate gels were run with the same set of protein samples and (a) one gel was stained with Coomassie blue. The other gel was transferred to a PVDF membrane, and (b) transferred protein was detected by staining the membrane with Azure Ponceau. (c) After destaining with water, the membrane in (b) was subjected to Western blotting to detect the 72-KDa protein of interest. Lanes 1-3, solubilized inclusion bodies. Lanes 4-6, insoluble pellets. The Western blot was detected using Radiance HRP substrate.

QUICK TIP:  While fine for PVDF and nitrocellulose membranes, Ponceau S will permanently stain nylon membranes. If you see poor transfer across the length of your lane, repeat the above advice: try a longer, cooler transfer, and remake your buffers.

STEP 3: Check Your Protein's Molecular Weight

If your protein has either a high (<120 kDa) or low (>25 kDa) molecular weight then you should consider altering your transfer conditions. For large proteins consider increasing the transfer time, sometimes even overnight, but keeping your transfer cool is key. For smaller proteins consider decreasing the transfer time and reducing the voltage to prevent proteins completely passing through your membrane.

The composition of your transfer buffer can have a huge impact on how readily your proteins move and bind.

For high molecular weight proteins, SDS can promote movement out of the gel and onto the membrane, whereas methanol promotes membrane binding but can inhibit transfer. Consider slightly reducing methanol to between 5-10% and adding SDS to a concentration of 0.1%.

For very small proteins which face the other problem, SDS should be eliminated from the transfer buffer, and methanol concentration increased up towards 20%.

A Ponceau S stained membrane showing efficient protein transfer. Transfer efficacy at higher and lower molecular weights is somewhat reduced.
A Ponceau S stained membrane showing efficient protein transfer. Transfer efficacy at higher and lower molecular weights is somewhat reduced.

QUICK TIP:  For very large proteins, it may be worth considering using a lower percentage gel to allow easier migration of proteins into the membrane. Conversely, small proteins readily leave the gel and pass into the membrane, to prevent transfer through check that your membrane has the smallest pore size of 0.2 μm.

Tried everything, and still no bands?

If at this point you’re transferring like a pro, but still aren’t seeing anything, keep reading. If you’re working with a very low-abundance protein or with a poorly validated antibody, at this point it is worth checking for something that should definitely be expressed. We’re here to help- fill out the form on this page to ask a Western blotting expert a question. Ruling out problems with your probing and developing is the next step; I’ll cover that in a later blog post.

Frequently Asked Questions

Ensure your sample buffer contains all of the ingredients needed, in the right concentrations, and that nothing has expired. If there is an issue with migration, it is probably best to make fresh sample buffer to be safe. Read more

Polyacrylamide gels form a crosslinked, mesh-like matrix through which proteins migrate with the application of current. The matrix functions like a sieve, allowing smaller, low molecular weight proteins to migrate quickly through the gel, while large, high molecular weight proteins tend to progress more slowly. When the electrophoresis is finished, proteins in each lane will be separated into a continuum, with the highest molecular weight proteins remaining near the loading site and the lowest molecular weight proteins reaching the opposite end of the gel. Read 6 tips on troubleshooting band separation

Gel documentation system use light from a wide variety of light sources and filters
to detect fluorescent dyes up to 832nm. contain an ultraviolet (UV) or visible (blue or white) transilluminator, a hood to block the samples from external light sources and a high-resolution camera for image capture.

QUANTITATIVE WESTERN BLOT BASICS
quantitative western blot basics

CLAIM YOUR FREE QUANTITATIVE WESTERN BLOTTING BASICS GUIDE

Get a quick overview of the steps you can take to ensure your Western blots are quantitative. This free guide also includes a troubleshooting section and tear-out quantitative Western blotting checklist.

Download Guide

Bendy bands and what to do with them

Categories
SDS-PAGE Transfers Troubleshooting

If you’ve worked with SDS-PAGE and Western blotting for a while, you’ll undoubtedly have seen bendy bands, or fuzzy bands, or both! This blog post will help you achieve those perfect razor-sharp rectangles shown on your antibody datasheet. While unattractive in their own right, these bendy bands make determining the molecular weight difficult and can interfere with band densitometry analysis, especially when using automated programs or when working with proteins that produce tight duplexes.

Fear not, here are five extremely simple steps you can take to cure the vast majority of band bending, and make your blots presentable again.

An example of some bendy bands in a precast gel from a ResearchGate post.
An example of some bendy bands in a precast gel from a ResearchGate post

Get Rid of Bendy Bands SDS-PAGE

STEP 1: Clean your gel running system

Picture this: you’ve spent weeks growing your rare cells, treated them with your incredibly expensive novel compound, carefully harvested and lysed your sample, only to cast your gel in between two dirty smeared plates. Yikes! To save yourself from this woe, always clean your plates and combs.

Azure Aqua during Western blotting experiment
The transfer cassettes and electrode core for the Azure Aqua Transfer Cell are colored for directionality which makes transferring easy and straightforward. You can perform a quick 1-hour transfer or you can transfer overnight at a lower voltage.
Electrophoresis Chambers

  • Quick Tip: Clean your plates and combs! Making sure they are spotlessly clean before casting ensures your perfect gels and perfect blots.

STEP 2: Shaken, stirred...

…and left to rest! A major issue with gel casting is improper mixing of reagents, in particular the acrylamide. In order to ensure a smooth separation of proteins across your resolving gel you want to ensure a constant acrylamide percentage throughout. But equally you want to get your gels cast and on with your experiment as quickly as possible. Here, as with many things in life, more haste equals less speed.

  • Quick Tip: A thorough mixing of gel reagents, followed by the equally important degassing step, will give you pristine gels first time, every time.

STEP 3: Fresh is best

We often have breaks in our Western blotting, and whilst it’s tempting to reach for that 6 month old bottle of running buffer, which has been sitting in the sun, it probably makes more sense to throw it out and start again. At the very least checking the pH of your stacking and resolving gel reagents is a must as this plays a fundamental role in how your proteins migrate through the gel.

  • Quick Tip: It’s worth keeping on top of your buffers and ensuring they’re fresh.

    Fresh buffers guaranties they have the correct pH.

Bendy bands SDS Page
Oddly shaped bands from ResearchGate

STEP 4: Slow and steady

We know you want to get your exciting results ASAP, but running your gels with a high voltage might leave you having to cast and re-run your samples again.

  • Quick Tip: Forcing your samples through the stack too quickly can mean they hit the resolving gel spread out, rather than in a nice tight line.

    The same is true in the resolving gel, by running your gels at too high a voltage the buffers and your gels become hot, and this is a major contributor to warping and bending in your gel.

STEP 5: More isn’t always better

Less about bending and more about blotting, this step is vital in ensuring you get the most accurate results possible from your blotting. While it’s tempting to load the maximum concentration of protein in the maximal volume, sometimes it’s worth reducing the load to ensure you get a nice sharp band rather than an unintelligible blot. 

  • Quick Tip: With new antibodies, and even new antibody batches,

    It’s always worth running a test with a standard curve of total protein. This simple step can save you sample, and time as it makes your analysis quicker and easier too.

If you’re looking for an imager to image your Western blots, your search ends here. Request a free, virtual demo of an Azure Imaging System, and say “Hello” to beautiful Western blots.

Scientist standing next to Azure 600
The Azure 600 gives you the flexibility you need for your research, while delivering solutions for quantitative Western Blot imaging. Azure Biosystems provides a unified Western Blot workflow, from the high-performance imaging system and analysis software to the reagents and consumables.
Browse Western Blotting Systems

Following these five steps will help you keep your bands razor sharp. They’ll be the pride of your lab-book, rather than something to leave tucked away in the back of your drawer. While you’re here, watch out for these assumptions about SDS-PAGE from our friends at NanoTemper.

Additional blog posts on SDS-PAGE:

FREE WESTERN BLOT eBOOK

New to Western blotting? Need to troubleshoot your Western blot?​ Want to brush up on Western blotting best practices? Claim your free Western Blotting eBook!
Download eBook

Linear range of detection and what it means for your quantifications

Categories
Multiplex Quantification Western Blotting

When quantifying Western blots many people assume it’s ok to simply image your protein of interest, strip, re-probe for their loading control of choice and then use this to normalize, without considering the linear range of detection. However, there are several pitfalls associated with this, and more and more journals are now asking for a proper defined normalization protocol, including evidence of linear detection, to be included with all manuscripts.

The linear range of detection is the region in which band intensity is proportional the amount of target protein on the membrane. As the amount of protein increases so should the band intensity. In the image below you can see where the issue lies, the linear range is marked with the blue box, outside of this region this proportional relationship is lost, especially at the extreme ends where the signal will be either completely over or under-saturated.

Linear ranges of detection overlap
Western blot quantification is only truly accurate if both the target protein, and loading control both have similar linear ranges.

Things get more complex still when loading controls are factored in as Western blot quantification is only truly accurate if both the target protein, and loading control both have similar linear ranges. This is shown in the two graphs on this below. In the graph above, the linear ranges of detection overlap; quantification of proteins within this overlap zone will be fine. However, in the latter where they don’t overlap it is easy to see how changes in protein concentration can impact the band-intensity for one protein, but not the other. In this instance quantification is next to useless.

Linear ranges of detection overlap
Western blot quantification is only truly accurate if both the target protein, and loading control both have similar linear ranges. This graph on the left shows linear ranges of detection overlap; quantification of proteins within this overlap zone will be fine. The graph on the left shows linear ranges that don’t overlap. It is easy to see how changes in protein concentration can impact the band-intensity for one protein, but not the other. In this instance quantification is next to useless.

Determining the linear range is relatively easy, and can be achieved by taking a sample and performing a serial dilution. If the ranges overlap then determining the amount of sample to load is also similarly easy. However, if the ranges don’t overlap, as is common when assessing low-abundance proteins in association with a highly expressed housekeeping gene, then it becomes more difficult to accurately quantify. Furthermore, if your protein of interest is strongly up or down-regulated by your treatment then this can further interfere with your attempts to fit within the linear range.

What to do if your linear ranges don't overlap

Don’t worry! There are several options available. Here are two tips on what to do if your linear ranges don’t overlap.

AZURE EXPERT TIP #1: Change loading control

This is an often-overlooked step. If you know your protein is expressed at relatively low levels then a loading control such as beta-actin will likely not be suitable. In these instances, using an organelle specific protein may prove more beneficial such as Lamin B1 or HSP60 as they should be less abundantly expressed, although care should be taken to confirm that expression of these is not altered by your treatments.

AZURE EXPERT TIP #2: Transition from loading controls towards total protein quantification

Transitioning from loading controls towards total protein quantification as a means of controlling for loading is the next thing you could try. By assessing the whole range of proteins over, or under-saturation becomes less of an issue generating a broader dynamic range. These stains include both in-gel and membrane stains and can be imaged in a variety of different ways. This technique is a great way to check both the quality of your protein separation and transfer efficiency.

You can also expand the dynamic range during imaging. Film has a notoriously poor dynamic range and issues with inaccurate exposure timings can further complicate matters. Gel and membrane imaging systems, such as our Azure Imaging Systems. Modern imaging systems such as the Azure 600 are able to image across a much wider dynamic range and therefore linear range of detection. Request a free demo of an Azure Imaging System, and say “Hello” to beautiful Western blots.

Two scientists working on Azure 600
The Azure 600 uses a 9.1MP camera to provide high resolution imaging perfect for publications. Change the sample to optics distance using adjustable height shelf for enhanced detection. Zoom into the area of interest with ROI imaging to reduce background.
Browse Western Blotting Systems

More posts on Western blotting:

Western blotting past, present and future

Categories
Western Blotting

We can’t think of a better way to start off our blog posts with a post covering the history of Western blotting and bust a few myths and misconceptions along the way.

Western blotting: fact or fiction?

Below are two statements regarding the history behind Western blotting. Can you guess which one is fact, and which is fiction?

Where did the name "Western blotting" come from?

So, if not Dr. Western, where does the name “Western blot” come from? To understand that we have to go look back at history.

In 1975, Edward Southern described his eponymous DNA transfer and probing technique, Southern blotting (R).

Two years later, in 1977, Stark reported on a new methodology to transfer and probe RNA, and as a simple pun named it Northern blotting (R). And so the groundwork was laid for Burnette to introduce another pun to the scientific world: the Western blot.

Since then, Western blotting has gone from strength to strength and is now a cornerstone protocol in life sciences. While there have been some refinements in buffers and materials, the general protocol remains remarkably similar to what was once used.

Western blot protocols

Western blot. Adapted from Renart J, Reiser J, Stark GR. PNAS, 1979.
Figure 1. One of the first Western blots. Adapted from Renart J, Reiser J, Stark GR. PNAS, 1979.

The future of Western blotting

After we’ve already covered the history of Western blotting, a question you might have is where next? The major ongoing development is the transition from classical film based exposure and imaging towards digital imaging systems. Delivering, higher levels of sensitivity across a wider dynamic range digital imagers are removing a lot of the hassle of Western blotting. But digital imagers aren’t just delivering improvements to existing techniques, the high-performance optics together with novel antibody conjugates and excitation devices are introducing new ways of performing and analyzing Western blots.

In-cell Western blotting: a new way to analyze Western blots

Fluorescently conjugated antibodies coupled with specific emission sources and filter sets are bringing Western blot multiplexing into the mainstream, stripping and re-probing, may become a thing of the past. However, while fluorescent antibodies are an evolution the standard Western blot protocol, requiring the same initial time investment, a new development representing a revolution in protein analysis is currently being developed.

In-cell western assay
In-cell Western assay performed using AzureCyto In-cell Western Kit. A 96-well plate was imaged on the Azure Sapphire FL using the 532, 685, and 784 Standard Optical Modules at a resolution of 10µm with the focus set to +3.00mm.

In-cell Western blotting is a radical rethinking of the initial Western blotting protocol coupling the ability to accurately quantify intracellular proteins from Western blotting with the repeatability, quick turnaround and high-throughput of an ELISA.

With in-cell Westerns, cells are grown on sterile plates, before being fixed and permeabilized in situ. Subsequent labelling is performed as normal with a specific primary antibody followed by an isotype specific near infrared (NIR) conjugated secondary antibody, to reduce auto-fluorescence and noise typically associated with tissue culture plastics. This ability to accurately investigate protein expression in situ is game changing drastically reducing time to result whilst also significantly expanding throughput.

When to use ELISA vs. Western blotting

As you can see, Western blotting as a technique is developing rapidly on several fronts. While new techniques may require slightly higher upfront costs; significant future savings in cost and time can be made. Furthermore, the increased selectivity and sensitivity these techniques deliver allow for more accurate quantifications from smaller sample sizes.

More posts on Western blotting: