Document

Western Blot Ebook

WESTERN BLOT EBOOK

TABLE OF CONTENTS

PAGE 5 Introduction to Western Blotting

PAGE 21 Imaging Western Blots

PAGE 31 Western Blotting Tips and Guidelines

PAGE 45 Protocols

PAGE 65 Application Notes

PAGE 91

WesternDot® 585

Selected PublicatioWWneessstteerrunnDDsooitt®®s68n5050g

Azure Biosystem Products

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ABOUT AZURE BIOSYSTEMS
At Azure Biosystems, we develop easy-to-use, high-performance imaging systems and high-quality reagents for life science research. By bringing a fresh approach to instrument design, technology, and user interface, we move past incremental improvements and go straight to innovations that substantially advance what a scientist can do. By focusing on getting the highest quality data from these instruments– low backgrounds, sensitive detection, robust quantitation–we’ve created a line of reagents that consistently delivers reproducible results and streamlines workflows.
Providing scientists around the globe with high-caliber products for life science research, Azure Biosystems’ innovations open the door to boundless scientific insights.
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A WORD ON QUANTITATION
MORE THAN JUST MEASUREMENT: WHY WESTERN BLOTS SHOULD ALWAYS BE QUANTITATIVE
Why start a guidebook on Western blotting with a discussion on quantitation? For many researchers new to Western blotting, the ease with which you can use an imager or scanner to quantify a band on a blot can sometimes lead to a scientist’s worst nightmare–non-reproducible results and incorrect conclusions. Just because you can generate a number doesn’t mean it’s true. But it also doesn’t mean that the number is never true.
So can I get quantitative data from Western blotting? Should I care even if I don’t want to quantify the bands on my blot? The answer to both questions is a resounding yes, and a review of what we mean when we use the word quantitative explains why.
What does quantitative mean, really? It can be surprisingly difficult to find a definition of the word quantitative that goes beyond a dictionary’s entry of “can be measured.” For scientists, it’s a word that’s used so often that the implication is everyone should know what it means. But different people use the word in different ways. At Azure Biosystems, we embrace the definition put forward by the statistician Samuel S. Wilks1, who said that to quantify something you must measure it, and your measurements need to meet the following criteria: · Your measurements are generated using a clearly defined process (the Western blot) · The measurement process generates a reproducible outcome, i.e the measurements are precise2 · The measurement process generates a valid outcome that reflects the “true” measurement, i.e. the measurements
are accurate3 The definition is a simple re-statement of good scientific measurement principles and helps remind us of what we need to achieve when we talk about generating quantitative Western blot data, i.e. Western blots that are reproducible and where we can verify measurement accuracy. When looked at in this way you can see that these goals are clearly within reach for Western blotting if enough care is taken to reduce or control for variability during the Western blotting process. Furthermore, it should be clear that whether you want a numerical value for the intensity of a band or you merely want to know if the band is there, you still want results that are reproducible. Therefore, every Western blot should be quantitative.
Making every Western blot quantitative By focusing on best practices for Western blotting and how to choose appropriate experimental conditions, this guidebook provides an excellent starting point and resource for researchers interested in generating reliable, reproducible Western blots. Whether your final data is numbers on a graph or simply the image of the blot, we hope we’ve helped you make all your Western blots quantitative.
References
1. SS Wilks. Some Aspects of Quantification in Science. Isis. June 1961. 52:2. p. 135. DOI: 10.1086/349466 2. Joint Committee for Guides in Metrology (JCGM). International vocabulary of metrology ­ Basic and general concepts and associated terms
(VIM). JCGM 200:2012, 3rd Edition 2012. p22. https://www.bipm.org/en/publications/guides/vim.html 3. Joint Committee for Guides in Metrology (JCGM). International vocabulary of metrology ­ Basic and general concepts and associated terms
(VIM). JCGM 200:2012, 3rd Edition 2012. p21. https://www.bipm.org/en/publications/guides/vim.html
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TABLE OF CONTENTS

1. Introduction to Western Blotting

5

1.1. Background ………………………………………………………………………………………………………………………………………………………………………………………….6

1.2. Gel Electrophoresis ……………………………………………………………………………………………………………………………………………………………………………..6

1.3 Transfer to Membrane ………………………………………………………………………………………………………………………………………………………………………..9

1.4. Membrane Blocking…………………………………………………………………………………………………………………………………………………………………………. 12

1.5. Membrane Incubation with Antibody …………………………………………………………………………………………………………………………………………… 13

1.6. Antibody Detection………………………………………………………………………………………………………………………………………………………………………….. 16

1.7. References ………………………………………………………………………………………………………………………………………………………………………………………… 19

2. Imaging Western Blots

21

2.1. Leaving the Darkroom ­ Moving to Digital Imaging for Better Quantitation………………………………………………………………………….. 22

2.2. Choosing a System ­ CCD Imagers, Scanners, and Hybrid CCD/Scanning Systems ……………………………………………………………. 24

2.3. Imaging Beyond the Western Blot ………………………………………………………………………………………………………………………………………………… 28

3. Western Blotting Tips and Guidelines

31

3.1. Tips for Successful Western Blot Transfer …………………………………………………………………………………………………………………………………… 32

3.2. How to Improve Your Chemiluminescent Western Blots …………………………………………………………………………………………………………… 34

3.3. How to Improve Your Fluorescent Western Blots……………………………………………………………………………………………………………………….. 36

3.4. Transition from Chemiluminescent to Fluorescent Western Blots……………………………………………………………………………………………. 37

3.5. Western Blot Normalization …………………………………………………………………………………………………………………………………………………………… 40

3.6. Troubleshooting ……………………………………………………………………………………………………………………………………………………………………………….. 42

4. Protocols

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4.1. HRP/Chemiluminescent Blot Detection with Radiance Substrate …………………………………………………………………………………………… 46

4.2. Fluorescent Western Blot Protocol with AzureSpectra Reagents ……………………………………………………………………………………………. 51

4.3. Azure HRP Stripping Buffer …………………………………………………………………………………………………………………………………………………………… 58

4.4. AzureRed Fluorescent Protein Stain……………………………………………………………………………………………………………………………………………… 59

5. Application Notes

65

Three-Color Western Blots with AzureSpectra Antibodies………………………………………………………………………………………………………………….. 66

Phosphorylated protein detection is more efficient by fluorescent Western blot……………………………………………………………………………… 68

Imaging Three-color Western Blots with the Azure 600………………………………………………………………………………………………………………………. 71

Superior electrophoresis results with Lonza reagents and the Azure Imaging Systems…………………………………………………………………..73

DNA Dye Detection Limits using Azure Imaging Systems……………………………………………………………………………………………………………………. 77

Imaging Viral Load in Chicken Embryos………………………………………………………………………………………………………………………………………………….. 79

Phosphor Imaging with the Sapphire Biomolecular Imager ………………………………………………………………………………………………………………… 81

Detecting Proteins In-Situ with In-Cell Western Blotting……………………………………………………………………………………………………………………… 83

Increasing Assay Efficiency with Four-Color Detection ………………………………………………………………………………………………………………………… 85

Accurate Western Blot Normalization with AzureRed Fluorescent Protein Stain…………………………………………………………………………….. 87

6. Selected Publications Using Azure Biosystems Products

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1 INTRODUCTION TO WESTERN BLOTTING
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INTRODUCTION TO WESTERN BLOTTING
1.1. Background
Developed in the late 1970s and early 1980s, Western blotting is a widely used analytical technique that can identify one or more specific proteins in a sample containing a complex mixture of proteins1-3. The process originally consisted of gel electrophoresis to separate proteins by molecular weight, electrophoretic transfer to and immobilization of the proteins on a solid nitrocellulose membrane support, probing of the membrane with antibodies specific for the protein of interest, and detection of the bound antibody using radio-labeled staphylococcal Protein A followed by autoradiography for visualization. Today, the Western blot continues to be a popular assay for analyzing protein expression and, in its current form, the first three steps are nearly identical to the original protocol. However, many technological advances in the ensuing years have increased the power of the approach. These advances include: · Improvements in the detection process that enable highly sensitive detection of low-abundance proteins · The use of safer, non-radioisotopic detection methods · The development of quantitative assays · The ability to detect multiple proteins simultaneously · The use of sophisticated digital capture systems for easier detection and analysis. And, like most useful methodologies, Western blotting technology continues to evolve. To help scientists new to Western blotting understand the fundamentals of the technique, and to provide those already familiar with Western blotting with an overview of the current state-of-the-art technologies and techniques, we’ve developed this guidebook. In these pages we review current best practices for Western blotting as well as the underlying physics, chemistry, and biology of the method so that scientists can take full advantage of this long-used and still powerful technique.
1.2. Gel Electrophoresis
In the first step of a Western blot, proteins are physically separated from one another across a gel matrix in a process called gel electrophoresis (Figure 1.1). A protein sample is mixed with a loading buffer, loaded onto the gel, and then subjected to an electrical current which is applied to the gel/buffer system. The proteins, which are negatively charged under the experimental conditions, travel through the gel towards the positive electrode. Depending on the type of gel and buffer system used, the distance a protein migrates through the gel matrix is governed primarily by the mass:charge ratio of the individual protein or simply the molecular weight of the protein. Protein electrophoresis can be run under either native or denaturing conditions (Table 1.1). Denaturing conditions are suitable for most applications. However, if the three-dimensional structure of the protein needs to be retained, native electrophoresis conditions must be used.
Figure 1.1. Polyacrylamide gel electrophoresis.
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Native

Features

Uses

· Proteins retain threedimensional structure and folding
· Proteins are separated by mass:charge ratio and cross-sectional area of proteins (Stokes radius)

· Use to study protein complexes
· Use when detection antibody recognizes an epitope on the folded protein

Denaturing

· Proteins are denatured using SDS (Figure 1.2) and heat
· Proteins are separated primarily by molecular weight

· Use when native conditions are not required

Table 1.1. Native vs. denaturing gel electrophoresis.

How To
· Omit SDS from loading buffer, gel and running buffer
· Omit denaturing agent from sample buffer
· Do not heat sample prior to loading
· Mix protein with sample buffer containing SDS and a denaturing agent
· Heat samples prior to loading

Gel Composition
The most commonly used protein electrophoresis approach is SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis). SDS-PAGE gels are composed of a pH-buffered solution, a mixture of acrylamide and bisacrylamide (the gel matrix components), and SDS. Ammonium persulfate (APS) and tetramethylethylenediamine (TEMED) are used to catalyze polymerization of acrylamide monomers and incorporation of bis-acrylamide ensures crosslinking of individual strands of acrylamide polymers.
The resolving capability of the gel is determined by the gel pore size, which is governed by both the concentration of acrylamide as well as the concentration of the bis-acrylamide crosslinker. In general, higher percentage gels have a smaller pore size and are used to separate proteins with lower molecular weights. Lower percentage gels have a larger pore size and are used to separate higher molecular weight proteins. Table 1.2 illustrates the resolving power of gels with different percentages of acrylamide.

Folded protein

SDS

SDS binds to the protein, resulting in denaturation and a uniform negative charge.
Figure 1.2. Mechanism of SDS denaturation of proteins.

MW Range, kDa

Gel Percentage

10­43

15

12­60

12

20­80

10

30­95

8

50­200

6

Table 1.2. Resolving power of different concentrations of acrylamide.

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Laemmli Discontinuous Gels
Discontinuous gels (also known as Laemmli gels) are among the most commonly used gel systems and can provide sharper, more defined bands than a continuous gel system4.
A discontinuous gel system consists of two stacked layers of gels, each with a different acrylamide concentration and pH, and a buffer that is at a different pH from both gels. The discontinuity between the pH of the two gels and the buffer alters the mobility of ions–specifically zwitterions–through the two gels, which in turn affects the mobility of the migrating proteins.
The top gel layer is called the stacking or focusing gel. The stacking gel contains a fixed low percentage of acrylamide and the lowest of the three pH levels. In a typical Tris-glycine buffer system, the pH for a stacking gel is 6.8, with a buffer at a pH 8.3. The purpose of the stacking gel is to concentrate all proteins into a single, tight band before they enter the lower portion of the gel, enhancing the sharpness and definition of individual protein bands in the resolving gel.
The lower gel, or resolving gel, contains the percentage of acrylamide needed to resolve the protein of interest. The pH of a typical resolving gel in a Tris-glycine buffer system is 8.8–more alkaline than both stacking gel and buffer. The purpose of the resolving gel is to separate proteins by size, and thus the composition of the gel (pH, percentage of acrylamide) is chosen to ensure that proteins move through the gel primarily based on molecular weight.

Gradient Gels
The resolving power of a gel can also be improved by using a gradient of acrylamide that increases in concentration from the top of the gel to the bottom, thus creating a “gradient gel.” In a gradient gel, proteins progress through the gel until the pore size impedes further migration.
Gradient gels are a great choice when you want to use a single gel to resolve multiple proteins that span a wide range of molecular weights.

Buffer Systems
In addition to varying the gel composition and setup, altering the electrophoresis buffer system can also optimize protein separation during gel electrophoresis (Table 1.3).
Standard SDS-PAGE gels use the alkaline buffer Tris-glycine, which provides adequate resolution for mid-size proteins. However, because the high alkalinity of the gel can lead to protein degradation and, thus, smearing of protein bands during longer run times, Tris-glycine is not ideal for resolving larger proteins (>150-200 kDa). Smaller proteins are also not easily resolved using Tris-glycine gels due to intermingling of SDS with the small molecular weight proteins in the stacking gel. This intermingling leads to fuzzy bands and reduced resolution of small (<10-15 kDa) proteins.
An alternative to Tris-glycine is the acidic Bis-Tris gel buffer that may be used in conjuction with two different running buffers: either MES (2-[N-morpholino] ethanesulfonic acid) buffer for small proteins or MOPS (3-[N-morpholino] propanesulfonic acid) buffer for mid-sized proteins. This system contains an additional reducing agent, sodium bisulfite, in the running buffer which works with the acidic Bis-Tris gel buffer to increase resolution and the sharpness of the protein bands.

Buffer System pH

Advantages

Tris-glycine

Up to 9.5

Good for mid-ranged proteins; inexpensive

Tris-Bis

6.4

Sharp protein bands; two running buffer options for optimization based on MW

Tris-acetate

7.0

High resolution of large MW proteins

Tris-tricine

Up to 9.5

Tricine separates low MW proteins from free SDS

Table 1.3. Buffer systems for protein gel electrophoresis.

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For resolving large molecular weight proteins, Tris-acetate gels are frequently used. The pH of Tris-acetate gels is close to 7.0, which supports protein stability during the long run times needed to adequately separate large molecular weight proteins. Tris-acetate can be used in both native and denaturing gels.
For resolving small proteins in the 1-10 kDa range, a Tris-tricine system is recommended. With a Tris-tricine gel and running buffer, free SDS can be separated from the low molecular weight proteins running near the leading edge of the gel front during electrophoresis.
1.3. Transfer to Membrane
After electrophoretic separation of proteins through the gel, the proteins are transferred to a solid membrane support for subsequent steps. Efficient transfer relies on the choice of membrane, the type of transfer apparatus used, and the composition of the transfer buffer.

Nitrocellulose Versus PVDF Membranes

Two types of membranes are commonly used for Western blots: nitrocellulose and polyvinylidene difluoride (PVDF). Both membrane types work well, but differences in experimental setup and sample composition will affect the choice of membrane. It should be noted that the signal intensity and background a sample produces on one type of membrane may be quite different on another type of membrane, even if the antibody and detection chemistry are the same.

Nitrocellulose membranes bind proteins strongly and nearly irreversibly. They are more cost effective than PVDF and were the first type of membrane used for Western blotting. Nitrocellulose is known for low background, particularly when using chemiluminescent detection (see below). Unlike PVDF, nitrocellulose does not require an activation step prior to use. However, nitrocellulose is fragile, making it difficult to manipulate the membrane, especially if the membrane is stored over a long period of time. Nitrocellulose also cannot be used with fluorescent detection methods as the membrane autofluoresces.

Like nitrocellulose, PVDF membranes have a strong affinity for proteins and are well-suited to Western blotting. They are more durable than nitrocellulose membranes making them popular for experiments that require multiple manipulations of the membrane. PVDF membranes are used for chemiluminescent and fluorescent detection and require incubation in methanol for activation prior to use. Low-fluorescence PVDF membranes provide the lowest fluorescent background and are recommended for fluorescent detection.

Conveniently cut, high-quality PVDF and nitrocellulose membranes are available from Azure Biosystems. To see these and all of our Western blotting reagents and consumables, visit azurebiosystems.com/reagents.

Transfer Conditions
Efficient transfer of proteins relies on both the migration of proteins out of the gel and retention of proteins on the membrane. Like gel electrophoresis, the transfer step uses electricity to move negatively charged proteins towards the positively charged electrode. Transfer efficiency is affected by the type of transfer apparatus used, the individual protein, the transfer buffer, and the transfer conditions.

Learn more about how to choose between wet and semi-dry transfer methods on page 32 in our Western Blot Tips and Guidelines chapter.

Wet or Semi-dry Transfer
Two types of transfer setups are popular for Western blotting, wet transfer (Figure 1.3 and Table 1.4) and semi-dry transfer (Figure 1.4 and Table 1.5).

Wet Transfer–In a wet transfer setup, a “transfer stack” is assembled, consisting of the gel and membrane surrounded on both sides by several layers of filter paper. The filter paper-gel-membrane-filter paper sandwich is surrounded by sponge-like fibrous pads on each side and this transfer stack placed into a cassette or holder.

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Prior to assembly, the transfer stack components are pre-wetted with transfer buffer. After assembly, the transfer stack is submerged in a tank filled with transfer buffer, and an electrical current is passed through the buffer to transfer proteins from the gel to the membrane.

Protein transfer from gel to membrane can be optimized by altering the buffer system (see next section), transfer time, and transfer voltage.

The wet transfer method is used for most blotting applications and delivers highly consistent results. The method is especially useful for transferring proteins that span a wide range of molecular weights and, because of its consistency, wet transfer is recommended when quantitative analysis will be performed.

One of the disadvantages of wet transfer is the requirement for a large quantity of buffer for each transfer. Another disadvantage is heat generation, especially during longer transfers. Excess heat can lead to inconsistent transfer across the gel, protein denaturation, and even breakdown of the gel. Thus, keeping the transfer apparatus cool by using ice packs, a cooling circulator, and/or by performing the transfer in a cold room is highly recommended.

Figure 1.3. A typical wet transfer setup.

Advantages

Disadvantages

· Flexibility: · Transfer conditions are easily adjusted · Multiple transfer buffer options enable more ways to optimize transfer
· Supports transfer of a broad molecular weight range at one time
· Compatible with extended transfer times for large molecular-weight proteins
· Can be used for quantitative Westerns

· Heating of buffer can interfere with transfer · Cooling mechanism and/or cold room space required
during transfer · Large volumes of transfer buffer are required

Table 1.4. Advantages and disadvantages of wet transfer.

Semi-dry Transfer–The semi-dry transfer setup uses a similar transfer stack to the wet transfer setup, but instead of getting submerged in a tank filled with buffer the transfer stack is placed directly between two electrode plates (Figure 1.4). Thus, this method avoids the large amounts of buffer needed in a wet transfer system (Table 1.5).

The big advantage of the semi-dry transfer setup is the fast transfer speed. Because the distance between the electrodes is minimized, a strong electrical field is generated which leads to rapid transfer.

Another advantage of this method is that two different transfer buffers can be used on each side of the transfer stack ­ one on the gel side designed to promote migration of proteins out of the gel and another on the membrane side that promotes retention of proteins on the membrane.

Electrode Plate

+

+

Electrode Plate

­

­

Power Cord
Figure 1.4. A typical semi-dry transfer setup.

Apparatus Cover Transfer “stack”
Apparatus Bottom

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However, the strong electrical field is also a disadvantage for this method. Low molecular weight proteins can be transported too far and move beyond the membrane. In addition, the longer transfer times needed for larger molecular weight proteins cannot be achieved because of the limited buffering capacity

Advantages

Disadvantages

· Transfer is rapid · Can use discontinuous buffer system to optimize
transfer of proteins · Little buffer is required · Easy to set up · Good for performing large numbers of blots analyzing
the same protein

· High intensity field strength may cause low molecular weight proteins to migrate through membrane
· Difficulty in transferring high (>120 kDa) molecular weight proteins
· Not recommended for quantitative Westerns
· Gel can dry out

Table 1.5. Advantages and disadvantages of semi-dry transfer.

Transfer Buffers
Transfer buffers contain several components to promote protein transfer (Table 1.6). Buffer components are optimized based on the type of transfer system being used (wet or semi-dry), the type of gel employed, the choice of membrane, and the protein of interest.
Buffer Properties–Transfer buffers must have strong buffering capacities to maintain conductivity and pH during transfer. The buffer pH must be different from the isoelectric point (pI) of the protein-of-interest (the pH at which the protein has a net charge of zero) or the transfer will not occur.
The most common buffer choice is Tris at a pH of 8.1-8.5, which is higher than the pI of most proteins. Higher pH buffers such as CAPS or carbonate can also be used and are recommended when transferring high molecular weight or basic proteins.
For extremely basic proteins, acetic acid is the recommended transfer buffer.
IMPORTANT: When using acetic acid as a transfer buffer, proteins become positively charged and migrate towards the anode, requiring that the orientation of the transfer stack be reversed.
Alcohol–Methanol (and sometimes ethanol) is added to transfer buffers to counteract the effects of SDS, which reduces protein binding to nitrocellulose membranes. Alcohol removes SDS from proteins thereby promoting protein retention on the nitrocellulose membrane.
However, alcohol can also induce precipitation of proteins in the gel, and cause basic proteins to become positively charged or neutral which inhibits protein migration out of the gel. Thus, the amount of alcohol used in the transfer buffer should be optimized for the protein of interest.
Only high quality, analytical grade alcohol should be used when preparing transfer buffers.
Note that the addition of alcohol to the transfer buffer is only required if SDS and nitrocellulose membranes are used. If SDS is not used, such as for running a native protein gel, or when PVDF membranes are used, then alcohol is not needed in the transfer buffer.
SDS–Although SDS can inhibit binding of proteins to nitrocellulose membranes, it promotes elution of proteins out of the gel by increasing protein solubility. However, SDS increases the intensity of the transfer and may alter the antigenicity of some proteins. When used, the concentration of SDS in the buffer should be titrated for each protein of interest but should never exceed 0.05%.
PVDF membranes should be used when SDS is included in the transfer buffer since SDS inhibits binding of proteins to nitrocellulose membranes.

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Component Buffer

Example Tris, CAPS, Carbonate

Alcohol

Methanol, ethanol

Detergent

SDS

Tris-tricine

Up to 9.5

Table 1.6. Transfer buffer components.

Use Support conduction, maintain appropriate pH Increase binding of proteins to membrane Promote migration of proteins out of gel Tricine separates low MW proteins from free SDS

Discontinuous Transfer Buffers–One of the advantages of a semi-dry transfer setup is the ability to use different transfer buffers on each side of the transfer stack, i.e. a discontinuous transfer buffer system. Discontinuous transfer buffer systems enable an additional level of transfer optimization–a buffer designed to promote protein elution from the gel can be used on the gel-side of the transfer stack and a different buffer designed to promote retention of proteins on the membrane can be used on the membrane-side of the transfer stack.

Voltage and Time
Transfer times and voltage settings should be optimized for each transfer. While proteins generally transfer more rapidly at higher voltages, transfer efficiency is not always consistent. Insufficient current and/or time may result in incomplete transfer, while high current and/or lengthy transfer times may result in loss of proteins via transfer through the membrane without retention. Transfer condition guidelines are usually supplied with the transfer apparatus. However, these conditions can be adjusted based on the protein of interest.
In general, longer transfer times, with a lower current, are used in wet transfer while short transfer times with high current are used with semi-dry systems.

1.4. Membrane Blocking
While the antibodies used in Western blotting typically have a high affinity for a specific protein, they also tend to bind non-specifically and with low affinity to the Western blotting membrane. This non-specific binding can result in a high background signal when the blot is imaged, reducing detection sensitivity. To keep background signal as low as possible, the membrane is incubated in a blocking solution after transfer. The blocking solution works by binding to the non-specific antibody binding sites on the membrane, thus preventing antibody binding through occlusion.
Optimizing blocking conditions is important for obtaining high-quality Western blot data, especially when quantitative information is desired. Insufficient blocking can lead to high background, reducing the ability to detect a specific signal. In contrast, excessive blocking can mask legitimate epitopes, and some blocking agents can interfere with detection reagents. Several different types of blocking agents are available, and the blocking solution should be optimized for each antibody:antigen interaction.
Blocking Buffer Components
Buffered Salt Solution
Blocking buffers contain a buffered salt solution compatible with the detecting antibody and method of detection. Generally, Tris- or phosphate-buffered saline (TBS; PBS) is used. However, PBS can interfere with alkaline phosphatase-based detection methods and phospho-specific antibodies. TBS should be the solution of choice when using these reagents.
Detergents
Low concentrations of mild, non-ionic detergents, such as Tween-20, make good blocking agents and can be added to the buffered salt solution. Detergents should be added just prior to use to prevent microbial growth, and care should be taken to make sure the detergent is completely solubilized to prevent artifacts, such as a nonuniform signal or background signal. Detergent should not be used as a blocking agent when fluorescent detection methods are used since some detergents autofluoresce and will cause high background signal.

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Protein Blocking Agents Protein blocking agents are very common and highly efficient, as nitrocellulose and PVDF membranes have a high affinity for all proteins. However, some proteins can interfere with the antibody:antigen recognition process or inhibit detection methods. Thus, the best blocking agent should be determined empirically for each experiment.
Non-fat Dry Milk–Non-fat dry milk is one of the most popular blocking agents. It is economical, easy to prepare and contains a mixture of proteins that are efficient blockers. However, milk contains the abundant phospho-protein casein, which can interfere with phospho-specific antibodies. In addition, biotin is a component of milk that can inhibit detection methods that rely on streptavidin/avidin.
Serum–Whole serum, usually derived from horse or fetal calf, is another blocker that contains a mixture of proteins. Serum is less commonly used for Western blots than other blocking agents and contains immunoglobulins that can interact with primary or secondary antibodies leading to high background.
Bovine Serum Albumin (BSA)–The albumin protein isolated from cow serum is often used in place of milk, particularly when using phospho-specific antibodies. However, some preparations of BSA contain tyrosine-phosphorylated proteins that can interfere with the assay if anti-phophotyrosine antibodies are used. BSA also cannot be used with anti-lectin antibodies as these antibodies will bind to the carbohydrates in BSA, causing high non-specific background.
Other Single Proteins–Other purified single proteins, such as casein protein, are available as blocking agents. The use of single proteins can help prevent cross-reactivity that can occur when milk or whole serum is used. Different proteins can be tested to optimize the blocking step.
Protein-free Blocking Agents Protein blockers can have cross-reactivity with primary antibodies, interfere with detection reagents and/ or mask epitopes. Protein-free blocking agents have been developed to alleviate these problems. The watersoluble polymer polyvinylpyrrolidone, typically used as a blocking agent in Southern blots, can also be used for Western blots. Alternatively, protein-free blocking agents containing proprietary formulas are available from multiple companies.
Blocking Conditions Blocking efficiency is dependent on time and temperature. For most experiments, one hour of blocking at room temperature is sufficient. However, if lengthier blocking times are used (>2 hours) then blocking should be performed at 4°C to prevent microbial growth.
1.5. Membrane Incubation with Antibody
After blocking, the membrane is ready to be probed with antibody and unbound antibody washed away. The factors that influence probing and washing include whether a direct or an indirect detection method will be used, the quality and type of antibodies available, the number of antigens to be detected, the type of enzyme or tag that will be used for detection, and the incubation and wash conditions.
Direct vs Indirect Detection Direct Detection Method In the direct detection method, the detection label (enzyme or tag) is conjugated directly to the primary antibody that recognizes the antigen of interest (Figure 1.5). This is the simplest form of detection requiring the fewest number of steps. While direct detection is fast and straightforward, it is often less sensitive than indirect detection as there are fewer steps in the process where signal amplification can occur. Also, adding a label directly to a primary antibody can be cost- and/or resource-prohibitive.
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INTRODUCTION TO WESTERN BLOTTING

Indirect Detection Method
The indirect detection method is more popular and often more sensitive than direct direction. With indirect detection, the antibody that recognizes the antigen of interest, called the primary antibody, is unlabeled (Figure 1.5). Detection happens through the addition of a labeled secondary antibody that recognizes a specific epitope on the primary antibody. This epitope is usually species-specific and is present in all antibodies generated in that species; for example several different primary antibodies generated in rabbits can all be recognized by the same anti-rabbit secondary antibody. Because of the broad specificity of the secondary antibody, one labeling reaction can result in detection of a wide range of primary antibodies, making this method highly cost-effective.

Detector Molecule

Secondary Antibody

Detector Molecule

Primary Antibody Target Membrane

Primary Antibody Target Membrane

Indirect
Secondary antibody is labeled.
Figure 1.5. Indirect vs. direct detection.

Direct
Primary antibody is labeled.

The use of a secondary antibody for detection can increase senstivity versus direct detection of the primary antibody when multiple secondary antibodies bind to a single primary antibody. Flexibility is also increased because the type of label used for detection can be changed easily through the use of a different secondary antibody.

Fluorescent secondary antibodies from Azure Biosystems deliver superior signal-to-noise ratios. Learn more a azurebiosystems.com/antibodies.

Antibody Quality and Type
One of the major factors influencing the outcome of a Western blot is the quality of the antibody, especially the primary antibody. High-quality antibodies–antibodies that have high specificity, high binding affinity, and low background binding–increase the specificity and sensitivity of the assay. Both monoclonal and polyclonal primary antibodies can be used for a successful blot.

Polyclonal Versus Monoclonal Primary Antibodies

Polyclonal Antibodies–Polyclonal antibodies are made using an organism’s immune response to the antigen of interest. They are isolated directly from the serum of immunized animals, typically rabbits, goats, donkeys, or sheep, and thus are a complex mix of all the antibodies generated by the animal. During the immune response, an individual B cell generates an antibody that recognizes a specific epitope on the antigen, with different B cells generating antibodies that recognize different epitopes. Thus, a “single” polyclonal antibody actually contains a mix of individual antibodies that recognize different regions of the same antigen. Because the individual antibodies recognize different sites on the antigen, more than one antibody can bind to the antigen at the same time, making polyclonal antibodies more sensitive than monoclonal antibodies. However, this wide range of binding specificities can sometimes lead to a higher background signal with polyclonal antibodies.

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INTRODUCTION TO WESTERN BLOTTING
Monoclonal Antibodies–Monoclonal antibodies are collected from the supernatant of a hybridoma cell line–a cell line created by fusing a single antibody-synthesizing cell with a myeloma cell. These cell lines are usually of mouse or rat origin. Because a single antibody-producing cell, or clone, is used, the antibodies generated by the hybridoma cell line are identical to each other, and all recognize the same epitope on the antigen. Monoclonal antibodies are used when detecting specific forms of a protein, such as phosphorylation status, or when multiple primary antibodies are used at the same time. However, they can also be used for general detection of proteins. Monoclonal antibodies tend to give a less robust signal than polyclonal antibodies as there is only a single antibody binding to an individual protein.
Secondary Antibodies
Multiple types of secondary antibodies can be used for Western blotting. The type of secondary antibody used depends on the species and class of the primary antibody, the detection method, and any other considerations that might warrant the use of a specialized secondary antibody.
Species and Class Type of Primary Antibody–While monoclonal antibodies are usually derived from mouse or rat cells, polyclonal antibodies can be isolated from a variety of animals. The secondary antibody must be specific for the species from which the primary antibody was derived. Therefore, anti-mouse or rat secondary antibodies must be used with a monoclonal antibody while anti-rabbit secondary antibodies should be employed when the primary antibody is isolated from rabbits.
Antibodies can be divided into different classes and subtypes (also called isotypes), depending on the type of heavy and light chain molecules that make up the antibody. The secondary antibody needs to be matched to the primary antibody class and/or subtype for appropriate recognition. Secondary antibodies can be purchased that recognize all classes (i.e. all mouse IgG antibodies) or a specific class, such as mouse IgG2a. Secondary antibodies that recognize all antibody classes are best used in situations where the class or isotype of the primary antibody is not known (such as when working with polyclonal antibodies). Secondary antibodies with a specific isotype are used when the isotype of the primary antibody is known, and can increase the specificity of the assay. Secondary antibodies with a specific isotype are also useful for detecting multiple proteins simultaneously as each secondary antibody can have a different label.
Specialized Secondary Antibodies for Western Blots
Anti H+L–Anti heavy and light chain antibodies (H+L) are specific for both the heavy and light chain portions of the target protein. This type of secondary antibody is often used when the class of the primary antibody is unknown.
Light-chain Specific–Antibodies that recognize only the light chain portion of the antibody are typically used when Western blotting is being performed after an immunoprecipitation. The use of this antibody prevents recognition of the heavy chain of the precipitating antibody.
Antibody Purification Both primary and secondary antibodies can be purified to reduce background.
Affinity Purification–Affinity purification uses an affinity column–a column that contains the antibody’s epitope immobilized onto a solid support–to remove antibodies that do not bind to the desired epitope. Affinity-purified antibodies have increased specificity due to the presence of fewer offtarget binding antibodies.
Pre-adsorption–Pre-adsorption of antibodies can also be used to increase the specificity of binding and reduce cross-reactivity. For example, a rabbit anti-IgG antibody might be pre-adsorbed against mouse, rat, and goat IgG to prevent recognition of other species of IgG. Pre-adsorbed antibodies are highly specific and pre-adsorbed secondary antibodies are recommended for multiplex Western blotting, i.e. Western blots where multiple proteins are probed simultaneously.
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INTRODUCTION TO WESTERN BLOTTING

Antibody Conjugates
An important consideration in the choice of a primary (direct detection) or secondary (indirect detection) antibody is the type of label that is conjugated to the antibody. Detection antibodies can be labeled with enzymes (for chemiluminescent or fluorescent detection), fluorescent tags, or other small molecules that are used to amplify the signal (biotin).

Enzymatic Labels–Detection antibodies can be conjugated to enzymatic labels. The enzymes are then used to cleave products that produce light (chemiluminescence) or fluorescence for detection. The two most commonly used enzymes are alkaline phosphatase (AP) and horseradish peroxidase (HRP). Many products are available for use with enzymatic labels making them highly popular. Due to the kinetics of enzyme reactions, visualization of the signal must be timed appropriately to capture peak signal. The reaction can also saturate, limiting the dynamic range of protein concentrations that can be measured accurately.

Fluorescent Labels–Detection antibodies can also be labeled directly with a fluorescent tag, also called a fluorophore, and fluorescence measured using an imaging system. Upon excitation with light at the appropriate wavelength, the fluorophore emits light at a slightly longer wavelength. The emitted light is then captured and converted into a digital signal by an imager.
Unlike detection methods that rely on enzymes, the amount of light emitted from the fluorophore is consistent and directly proportional to the amount of protein on the membrane, making the assay truly quantitative. In addition, fluorescent labels are an excellent choice for multiplexing–simultaneous detection of multiple proteins in a single blot. For multiplexing, the different detection antibodies are labeled with fluorophores that emit light at distinct, spectrally separated wavelengths.

Other Conjugates–Biotin-labeled (biotinylated) detection antibodies can be used for sensitive detection of low-abundance proteins. This system relies on the strong binding interaction between biotin and fluorescently-labeled (or chemiluminescently-labeled) avidin/streptavidin. The detection antibody is typically conjugated to multiple biotin molecules, which results in a strong signal when multiple labeled avidin/streptavidin molecules bind to the biotin moiety.
1.6. Antibody Detection
Antibody binding can be visualized using colorimetric, chemiluminescent, or fluorescent detection methods. This guide focuses on chemiluminescent and fluorescent detection. The choice of detection method should be made based on multiple factors including the desired sensitivity.

Chemiluminescence

Chemiluminescence is a popular indirect detection method for Western blotting that relies on an enzymesubstrate reaction that emits light (Figure 1.6). Horseradish peroxidase (HRP) and alkaline phosphatase (AP) are two commonly used chemiluminescent enzymes, with the sensitivity of detection dependent on the choice of substrate–commercially available substrates for HRP can detect proteins in the femtogram range. Imaging of a chemiluminescent Western blot is historically done via exposure of the blot to x-ray film (Figure 1.7) and can also be done using a CCD-based imaging system.

Chemiluminescent detection is often used because it is specific, easy to perform, and highly sensitive–proteins can

be detected at femtogram levels (Table 1.7). The technique is very good at answering the question, “Is my protein

there or not?” however chemiluminescent detection is not very good at addressing questions such as, “How much

of my protein is present relative to another protein? How much of

my protein is in one sample compared to another sample? How do I control for sample loading inconsistencies?” We discuss the drawbacks to chemiluminescent detection in the next section.

Learn how to optimize your chemiluminescent Western blots on page 35 in our Western Blot Tips and Guidelines Chapter.

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HRP Secondary Antibody

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Primary Antibody

Protein Sample
Figure 1.6. Chemiluminescent Western blotting–one signal, one protein. In chemiluminescent detection, the antigen-primary antibody complex is bound by a secondary antibody conjugated to an enzyme, such as horseradish peroxidase (HRP). The enzyme catalyzes a reaction that generates light in the presence of a luminescent substrate, and the light can be detected either by exposure to x-ray film or by a CCD-based imaging system.

Figure 1.7. A chemiluminescent Western blot detected on x-ray film.

Chemiluminescence Drawbacks
Unlike fluorescent tags, where one or more different proteins can be probed simultaneously using antibodies labeled with spectrally distinct fluorophores, chemiluminescent reactions emit light over a broad range of wavelengths. Thus, with chemiluminescent detection, emission wavelengths cannot be used to distinguish signals from different proteins. Instead, the proteins must be well-resolved electrophoretically.
For example, proteins with small differences in molecular weight, such as the same protein with and without a posttranslational modification, tend to co-migrate during electrophoresis making them difficult to visualize simultaneously using chemiluminescence since the bands will most likely overlap.
Overlapping bands can also impact detection of normalization and loading controls. Unless these controls are wellresolved electrophoretically from the protein-of-interest, the blot must be either stripped and reprobed to detect the control, which renders the blot non-quantitative, or the controls must be placed on a separate blot, which is not a true loading control.
Furthermore, because chemiluminescence relies on an enzyme-substrate reaction, the amount of signal (emitted light) is subject to variations in reaction kinetics, variations which can be affected by reaction conditions, i.e. pH, temperature, substrate concentration, and enzyme concentration. This inherent variability makes chemiluminescence, at best, a semi-quantitative detection chemistry.
Finally, the traditional use of x-ray film as a method of visualization suffers from dynamic range limitations of the film that can often lead to signal saturation.

Advantages · Sensitive · Easy, familiar chemistry · Compatible with film or digital imaging
Table 1.7. Chemiluminescent detection.

Disadvantages
· Semi-quantitative · Signal dependent on enzyme kinetics · Single protein only, loading controls require stripping
and re-probing

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Fluorescence

Fluorescent Western blotting uses secondary antibodies directly conjugated to fluorescent dyes. Unlike chemiluminescent Westerns, which are limited by the variable kinetics of the enzyme-substrate reaction, the amount of light emitted from fluorophores is highly consistent and directly proportional to the amount of protein on the membrane. This consistency means that fluorescent detection can provide a truly quantitative analysis of the proteins in question.

Fluorophores for fluorescent Western blotting can be chosen based on their specific excitation and emission spectra, enabling multiplexing (detection of multiple proteins simultaneously, Figure 1.8) for faster, more efficient studies. Thus, one of

Find out how to improve your fluorescent Western blots on page 36 in our Western Blot Tips and Guidelines chapter.

the biggest advantages of fluorescent detection versus

chemiluminesence is the ability to use more than one antibody

per assay. With multiplexing, normalization and loading controls can be imaged at the same time and on the

same blot as the sample. In addition, the ability to use different fluorophores enables visualization of proteins that

are not well-separated electrophoretically, for more convenient imaging of the same protein with and without

post-translational modifications.

Secondary Antibody

Protein A

Primary Antibody

Secondary Antibody

Protein B

Primary Antibody

Figure 1.8. Fluorescent Multiplex Western Blotting. Multiplex detection is possible by using two or more fluorescent dyes and an instrument that can excite and detect the light from each dye.

Figure 1.9. Multiplex fluorescent Western.

Fluorescent Westerns are typically visualized using a digital imager rather than X-ray film (Figure 1.9). The newer generation of imaging systems often contain sophisticated cameras that exhibit a broader dynamic range than film, thus avoiding the signal saturation problems that limit the dynamic range of film. Finally, fluorescent dyes are relatively stable; blots can be archived and imaged months after the initial experiment as long as precautions are taken to avoid photo-bleaching of the fluorophores.
Fluorescence Drawbacks

Learn more about doing multiplex fluorescent Western blots in our application notes:
· Three-color Western bBlots with AzureSpectra Antibodies (page 66)
· Imaging Three-color Western Blots with the Azure 600 (page 71)
· Increasing Assay Efficiency with Four-Color Detection (page 85).

While fluorescent detection is the best choice for quantitation and can greatly accelerate workflows for analyzing multiple proteins via Western blotting, the method has a few drawbacks (Table 1.8). Fluorescent detection can be less sensitive than chemiluminescent detection, depending on the protein being assayed. Reagents (e.g. bromophenol blue) and supplies (e.g. certain membranes) can autofluoresce leading to high background which reduces the limit of detection of the assay. When switching from a chemiluminescent assay, all primary and secondary antibodies need to be titrated individually to find the highest signal-to-noise ratio. Lastly, fluorescent Western blots are visualized using digital imagers rather than the x-ray film and developer paradigm established with chemiluminescent detection.

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Advantages
· Multiplex capability · Increased quantitative accuracy · Fluorescent label stability allows blots to be stored and
re-imaged later
Table 1.8. Fluorescent detection.

Disadvantages
· Can be less sensitive than chemiluminescence · Membranes auto-fluorescence can increase background

Chemiluminescence vs Fluorescence

Chemiluminescent and Fluorescent Westerns: Choose the Best Assay for Your Experiment

Both chemiluminescent detection and fluorescent detection are excellent methods and, when used together, can provide complementary information that enhances insight. Thus, a laboratory should not be a “chemiluminescent Western laboratory” or a “fluorescent Western laboratory” but a lab that uses the best assay for each experiment. Table 1.9 outlines when to use chemiluminescent detection and when fluorescent detection might be best.

Find out how to transition from chemiluminescent to fluorescent Western blots on page 37 in our Western Blot Tips and Guidelines chapter.

Use Chemiluminescence to

Use Fluorescence to

· Detect a single protein · Assay for presence/absence of a protein · Measure antibody responses · Follow protein purification · Detect low abundance proteins

· Detect multiple proteins simultaneously · Study posttranslational modifications · Have same-blot loading control · Have in-lane normalization · Perform quantitative Westerns

Table 1.9. Chemiluminescent detection vs. Fluorescent detection.

1.7. References

1. Renart J, Reiser J, Stark GR: Transfer of proteins from gels to diazobenzyloxymethyl-paper and detection with antisera: a method for studying antibody specificity and antigen structure. (1979) PNAS 76:3116-3120.

2. Towbin H, Staehelin T, Gordon J: Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. (1979) PNAS 76:4350-4354.

3.

Burnette WN: Western blotting: electrophoretic transfer of proteins from sodium dodecyl sulfate-polyacrylamide gels to unmodified

nitrocellulose and radiographic detection with antibody and radioiodinated protein A. (1981) Analytical Biochemistry 112:195-203.

4. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. (1970) Nature 227 (5259): 680­685.

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2.1 Leaving the Darkroom ­ Moving to Digital Imaging for Better Quantitation
Digital imaging hardware and the associated quantification software are becoming ever more popular, surpassing film-based methods for gathering data. Compared to film, digital imaging offers a wider dynamic range, more accurate quantitation, and the ability to conduct multiplex imaging, all of which work together to greatly streamline workflows.
Here we discuss the key benefits of leaving the darkroom, and moving to digital imaging.
Detection of Signal Saturation
When capturing data from your Western blot, through film or a digital imager, you are measuring the intensity of signal collected in a specific area of the blot. When the signal accumulation for a specific band on a Western blot reaches the saturation level, no further signal accumulation is possible. This means that if two bands on your blot are saturated, you have no way of knowing if they have the same signal level, or if their signal level is vastly different.
Using film, detection of saturation is essentially guesswork and it is easy to miss variations in protein levels or to underestimate the amount of protein present.
Digital imaging has two advantages. First, software for the quantification of digital imaging can notify the scientist when the acquired image has bands that are saturated. (Figure 2.1 A, B) Second, software for digital imaging can determine the optimal image time that prevents signal saturation of the most intense band. This feature can prevent over-saturation of the signal, making sure the data is within the linear range of the imaging system.

A.

Film

Digital Image Showing saturation

B.

Sample Western blot

Same blot showing saturation
Figure 2.1. A wider dynamic range and automatic saturation detection allow for improved quantification. A. The same blot was imaged on both x-ray film and the Azure 600. The Azure 600 detects when saturation occurs and calculates an auto-exposure time to avoid saturated bands. B. The top blot is a sample western blot showing a variety of band intensities. Below is the same blot showing saturated bands, this blot should therefore be exposed for a shorter period of time for more accurate quantification, a process that is easy to do with today’s digital imagers.
Wide Dynamic Range
Dynamic range is the ratio of the maximum detectable signal to the background signal (i.e. noise). When comparing two different systems, the one with a wider dynamic range should provide faster workflows and more robust quantitation since both weak and strong signals can be measured during a single exposure. Without a wide dynamic range, strong signals will saturate before weak bands can be detected, requiring multiple exposures to visualize all bands and making quantitation either less reliable or even impossible (Figure 2.2).

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1 sec 10 sec

16-bit CCD system R2 = 0.99185

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12-bit CCD system
R2 = 0.75929

Log Pixel Intensity Log Pixel Intensity

0.10

1

10

100

0.10

1

10

100

Log Picograms Protein

Log Picograms Protein

Figure 2.2. 16-bit Imaging for a Wide Dynamic Range. A Western blot was imaged on both a 16-bit system and a 12-bit system. While the 10 second exposure appears similar on the different systems, the 12-bit system produces an image that is saturated and not suitable for analysis.

High Resolution
Whether you’re looking at a CCD camera­based system or a scanning imager, the resolution will determine the smallest feature size that you will be able to image. For CCD-based imagers, resolution is typically presented in megapixels (MPs), with a larger number of MPs providing a higher resolution. For scanning imagers resolution is provided in microns and indicates the smallest feature size you will be able to visualize. Higher resolution systems can facilitate imaging blots with poorly-resolved protein bands and can facilitate visualizing samples other than blots. For example, some high resolution scanning imagers can image individual cells within a tissue sample or microscope slide.

Multiplex Assays
Film and chemiluminescent imaging limit the assay to one signal detected per Western blot. The most recent generation of digital imagers have introduced the possibility of multiplex analysis (Figure 2.3) to assays that were previously only able to analyze one protein at a time. Fluorescent antibodies spanning the visible and near infrared spectrum enable detection of multiple samples of interest within a single assay, making within-experiment controls possible, supporting the development of novel assays, and greatly improving and accelerating workflows.

Red channel

Green channel

Merged image

Image Digitization
To include a Western blot image in any publication, digitization of the image is required. With film, scanning of the film is required in order to analyze or quantitate the image. With digital imaging, the resulting image is immediately digitized and ready for quantitative analysis with software.
To summarize, digital imagers represent a significant improvement over traditional film imaging with increased sensitivity, dynamic range and image quality. Furthermore, they offer significant quality of life improvements such as automatic saturation detection and quantification, while also opening the door to the development of new assays through their multiplex capacity.

Figure 2.3. Multiplex imaging significantly optimizes workflows and allows for new assay development. Multiplexing on a single blot is a unique method available with digital imagers. Here, a western blot has been probed for both STAT1 (red channel) and phosphorylated STAT1 (green channel) using fluorescently conjugated secondary antibodies and imaged using infrared detection on the Azure 600.

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2.2 Choosing a System ­ CCD Imagers, Scanners, and Hybrid CCD/Scanning Systems
Once a blot has been prepared the last step in the Western blotting workflow is image capture and analysis. Using the right imager and following best practices for data acquisition and analysis is just as important as choosing the right electrophoresis and blotting conditions when you want to generate reliable, high-quality Western blotting data. The choice of imaging system will depend on the type of studies being done, specifically the detection chemistry used and the sensitivity and resolution needed.

Different Imaging Systems for Different Performance Needs
Today’s digital imagers typically fall into one of two categories–CCD-based imagers and scanning systems. Choosing between the setups comes down to performance needs and whether you need the flexibility to support multiple imaging chemistries and applications. In both types of systems, one of the most important features to look for is dye flexibility. Not every fluorescent dye works with every imaging system, thus it’s important to verify that the system has enough dye flexibility that it will work with the dyes used in your studies. This is especially true when you want to do multiplex fluorescent Western blots. For multiplexing, it is critical to look at systems with multiple excitation sources and multiple detection wavelengths. Additionally, you will want to make sure that the best light sources have been selected. For example, IR dyes are typically excited with lasers, in part because only lasers emit light close to the excitation peak of these types of dyes.
Here we discuss the key features to use for imaging system evaluation.

CCD Imagers

CCD imagers use a CCD camera for signal detection. The blot (or gel, or other sample being imaged) is placed into a chamber and, if necessary, illuminated using a light source. The emitted light is then captured and digitized by the sensor. With this type of system, because the entire sample is imaged at once, the uniformity of sample illumination and sample detection is critical. It is therefore important to consider the imaging field of view (FOV) in CCD platforms, the larger the FOV the more difficult it is to control illumination uniformity. Excitation uniformity is easier to control with a smaller field of view.

Imaging chemiluminescent Western blots is most effectively done using a CCD sensor. The CCD sensor samples the entire light spectrum simultaneously and efficiently, supporting the long integration times (from seconds to minutes) needed for sensitive detection. Like film exposures, CCD imaging allows the user to control the exposure time.

Azure Biosystems offers CCD imaging systems that combine affordability with performance. Learn more about our Azure Imaging Systems at azurebiosystems.com/ imaging-systems.

Key Features for Evaluation of CCD Imagers
High Resolution, High Sensitivity–One feature essential for digital imaging with a CCD camera is pixel binning. The advantages of pixel binning–higher sensitivity and a higher signal-to-noise ratio–are the result of combining neighboring pixels into a single larger pixel, or “super-pixel” (Figure 2.4). The larger size of the super pixel increases sensitivity by increasing the surface area available for photon detection without similarly increasing the noise (note that the lack of increased noise only applies to on-chip binning and is not a feature of binning performed computationally during data analysis). For example, a binning of 2×2 combines four adjacent pixels into a single super-pixel resulting in a four-fold increase in sensitivity to light, while keeping the noise the same as that of a single unbinned pixel. While the increased pixel size can greatly improve the sensitivity of the detector, the larger pixel size reduces the resolution. A binning of 1×1 uses the native pixel size–no pixels are combined–and thus takes advantage of the full resolution of the sensor.

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1X1 Binning

5.4 µ 5.4 µ
10.8 µ

40 seconds, unbinned

2X2 Binning

10.8 µ

40 seconds, 2X2 bin

Figure 2.4. Pixel binning improves image sensitivity. Pixel binning is a powerful technique for digital imaging that increases sensitivity by combining pixels to make a larger “super-pixel.” The super-pixel has a higher signal-to-noise ratio (SNR) than the unbinned pixel.

It is important to understand that with high levels of binning the final image will have significantly lower resolution, and that low resolution images may not be suitable for publication. Upscaling, a form of post capture image manipulation, can be used to rescue the lost resolution, but an upscaled image must not be used for quantitative purposes because pixels (data) is artificially added to the image and only raw data should be quantified.
Wide Aperture for Chemiluminescent Detection–Especially for chemiluminescence, the F stop is an important value to consider. The smaller the F stop, the wider the aperture, and the more light can be let in. Imagers with a small F stop can deliver more light to the sensor, reducing exposure times.
Wide Range of Light Sources–While chemiluminescent does not require a light source, fluorescent imaging does require both a specific light source and emission filter.
There are two main light sources used in fluorescent imaging: light emitting diodes (LEDs), and lasers. Historically, LEDs have been used because of their wavelength flexibility and low cost. But laser technology has advanced rapidly and now offers similar wavelength flexibility at a competitive price. Lasers are fundamentally different from and have several advantages over LEDs. Because lasers are monochromatic, they are highly effective at exciting fluorophores at a precise wavelength. In a multiplex experiment, precise excitation is critically important for specific excitation to avoid bleed through. LEDs, which are a broad band light source, can excite many more fluorophores but may also introduce cross excitation / bleed through if proper care is not taken in the experimental design.
Scanning Systems
Scanning systems use laser light for sample excitation and either a photomultiplier tube (PMT) or avalanche photodiode (APD) to detect the emission signal. The sample is typically placed on a bed area and light excites the sample from the bottom. The emitted light can then be detected by a PMT or APD. Unlike CCD imagers, which image the blot or other sample all at once, scanning systems illuminate the sample one small section at a time as the sample is scanned. While scanning can take slightly longer than imaging with a camera-based system, the higher intensities possible with pinpoint rather than widely-distributed illumination can lead to greater sensitivity for more demanding studies. In addition, the ability to use PMTs and APDs, which are not compatible with a camera-based imaging approach, expands the range of detection chemistries and offers lower detection limits and overall better performance for visible and near infrared (NIR) fluorescence imaging.
Imaging fluorescent Western blots can also be done using a CCD sensor, but better performance can be obtained in a bed scanning system. Fluorescence imaging with a CCD imager is often less sensitive than chemiluminescence because fluorescence imaging does not enjoy the enzymatic signal amplification afforded by the horseradish peroxidase-coupled antibody. Fluorescence imaging with a laser scanner, however, can match or exceed chemiluminescence sensitivity because PMTs and APDs have an internal gain which serves to amplify the emission signal in a similar manner as the enzymatic amplification. Because CCD sensors cannot amplify the emission signal like a PMT or APD, laser scanners using a PMT and/or APD are the best choice for high sensitivity fluorescence imaging.

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Key Features for Evaluating of Scanning System

Resolution–Scanner resolution is important for application flexibility. Scanning systems enable fine tune control over the spatial resolution, allowing a user to select a resolution appropriate for a specific experiment. Most Western blots need only 100-200µm resolution, but some imaging application require much higher resolution to capture fine detail. For example, imaging stained tissue sections (IHC) may require 10µm resolution to visualize fine details in the tissue.

Scan Speed–Unlike CCD imagers that image the whole field of view at once, scanning systems scan the image line by line. The larger your scan area, the longer your scan will take. This is why it is important to select an imaging system with a fast scan speed, usually listed in cm/s. The faster the scan speed, the less time imaging your blot will take.

Excitation Sources–Scanner devices used for detecting fluorescence most commonly employ laser light for excitation. A laser is a collimated light source, meaning it produces a column of light such that nearly all the photons are travelling in the same direction/angle and nearly all the photons leaving the light source are effectively delivered to the endpoint (sample of interest/fluorophore).
By contrast LEDs, which have high divergence, produce a diffused pattern of excitation light where the light scatters/diffuses in many directions. When exciting a fluorophore, the more energy delivered to the fluorophore the greater the emission signal. Because lasers are a collimated light source and do not diverge like LEDs, lasers are much more effective at delivering excitation energy to the fluorophore and therefore produce a brighter emission signal yielding a lower limit of detection compared to LED excitation. This technological difference is critically important to achieving high quality images and data.
Another factor to consider is simply the number of light sources. The more lasers, the more dye flexibility. Furthermore, more lasers enable the detection of more fluorophores in a single sample ­ for example, a system with four unique lasers would enable the detection of four fluorophores from a single sample, increasing the multiplexing capacity, application flexibility, and throughput.

Detector Technology and Light Collection–Quantum efficiency and spectral response are important factors to review when evaluating scanning systems. It is also important to consider the number of detectors included in the scanning system. There are two main types of detectors used in scanning systems: Avalanche Photodiodes (APDs) and Photomultiplier Tubes (PMTs):

PMTs–PMTs belong to a class of vacuum tubes that convert photons into an electric signal. Photo multipliers have high internal gain, making them ideal for low light applications. The high internal gain is essential to high sensitivity fluorescence imaging.

Photomultiplier tubes operate using photoelectric effect and secondary emission. When light

is incident on the photocathode, it emits electrons into the vacuum tube. These electrons are

focused towards the electron multiplier (dynode), which multiplies the signal by secondary emission. The multiplied electrons

Focusing Secondary electrode electrons

are converted into an output signal by the

anode (Figure 2.5).

PMTs are ideal for low light imaging, where internal gain is required in order to amplify the signal. This is a key reason why they are used for phosphor imaging and other high sensitivity blue light detection ie GFP detection.

Primary electron Dynode

Anode

Photocathode
Figure 2.5. How a PMT works.

Connector pins

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APDs–An APD is a highly sensitive semiconductor electronic device that exploits the photo electric effect to convert light to electricity (Figure 2.6). APDs are similar to PMTs in that they also amplify the emission signal resulting in enhanced sensitivity through signal amplification. However, APDs are quite different from PMTs. The most important difference between APDs and PMTs is their quantum efficiency. APDs have a significantly higher quantum efficiency at longer wavelengths compared to PMTs, and because APDs can capture light more effectively than PMTs at longer wavelengths they are better suited for fluorescence imaging at wavelengths greater than 500nm . Combining the low autofluorescence associated with long wavelength fluorophores and the high quantum efficiency of APDs results in one of the most sensitive fluorescence detection systems. Although, bright blue dyes with a PMT detector can offer same or Figure 2.6. APD. similar performance.

Z-axis Flexibility–While all scanners can scan in the both X and Y direction, not all systems can freely scan in the z- axis. Controlling the z-axis scan plane allows users to easily image a variety of samples. Membranes, gels, culture plates, and slides all have different optimal z-axis focal planes, so it is important to consider a system than can easily change the z-axis for optimal imaging of a variety of samples.

Hybrid CCD/Scanning Systems

Hybrid CCD/scanning systems are a new class of imaging systems offering the best performance for a wide variety

of chemistry and sample types. As discussed above, there are CCD systems which are ideal for chemiluminescence

and gel imaging because they enable user control of the exposure time ­ short exposure for gel imaging and

longer exposures for high sensitivity chemiluminescence imaging. However, because CCD detectors do not

have a built in signal amplification mechanism they are not ideal for high sensitivity fluorescence imaging.

Alternatively, laser scanners using PMTs and APDs can amplify fluorescence signals and provide a highly sensitivity

fluorescence imaging system. Laser scanners however, do not allow for long exposures required for high sensitivity

chemiluminescence detection, so laser scanners are not ideal for chemiluminescence imaging. Combining the

advantages of a CCD imager for gels and chemi imaging with the

high performance of laser scanning system using PMT and APD detectors for high sensitivity fluorescence offers the best of both worlds. Hybrid CCD/scanning systems enable high performance across multiple imaging modalities and a wide variety of sample types for high application flexibility ­ this is the ideal imaging

Azure Biosystems offers a first-in-class hybrid CCD/scanning system, the Sapphire Biomolecular Imager. Learn more at azurebiosystems.com/sapphire.

system both chemiluminescent and fluorescent imaging.

CCD Imager vs Scanner vs Hybrid System: How to Choose
How to choose an imaging setup depends on the types of studies being done. For many labs, the performance of a CCD-based system covers most needs, especially for gels and chemiluminescent Western blotting. However, for labs that need high performance (i.e. sensitivity, resolution, dynamic range), to image large samples, and/or to use multiple imaging chemistries (phosphor imaging, chemiluminescence, and visible and NIR fluorescence) a scanning system may be a better choice.

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2.3. Imaging Beyond the Western Blot
Azure Biosystems offers CCD imagers and hybrid CCD/scanning systems that provide the value and versatility of imaging more than Western blots. Here are just a few examples of the different things you can image with our Azure Imaging System and Sapphire imagers.
Azure Imager System Images

Virulent Newcastle Virus Vs. “Normal” Newcastle Virus in chicken embryo imaged on Azure 600. Images provided by Ray Izquierdo-Lara, Ana Chumbe, Katherine Calderon, Manolo Fernandez. FARVET SAC, Peru.

Mouse imaging. Acquisition was in RGB automatic mode ­ red channel was imaged at 704 ms and green channel for 703 ms, both at 60 µm resolution. Imaged on Azure Imager System.

An agar plate with E. coli expressing GFPmut3 (green) and mCherry (red). The plate was imaged using red and green LEDs. Imaged on Azure Imager System.

Yeast colonies expressing GFP imaged in the Blue channel. Imaged on Azure Imager System.

Western blot using Cy5 and Cy3 of FtsZ1 and FtsZ2-1 from Arabidopsis thaliana. Plant fluorescent Western. Imaged on Azure Imager System.

GFP expressing cell monolayers with different protocols for fixation in different columns. Imaged on Azure Imager System.

2D Fluorescent Gel Imaged in Cy5/Cy3/ Cy2. Imaged on Azure Imager System.

Native Gel. Imaged on Azure Imager System.

TAMRA/GFP. Imaged on Azure Imager System.

GFP/mCherry. Imaged on Azure Imager System.
28

Antibody Array. Imaged on Azure Imager System.

Mouse with RFP-expressing subcutaneous tumor.

Sapphire Images

IMAGING WESTERN BLOTS

In-Gel Fluorescence and 2D Gels. Untreated and treated HeLa lysate were labeled with Cy3 and Cy5 respectively and simultaneously separated using IEF in the first dimension and SDS-PAGE in the second dimension then scanned at 100µm using the 520nm and 658nm lasers of the Sapphire Biomolecular Imager.

Rat Brain. Three color composite image of a rat brain scanned at 10µm on the Sapphire Biomolecular Imager.

Bee Head. Composite image of a bee head scanned with the 488nm and 785nm lasers of the Sapphire Biomolecular Imager.

Mixed Tissue. Composite image of mixed tissue scanned at 20µm with the 658nm and 785nm lasers of the Sapphire Biomolecular Imager.

Chicken Liver. Coomassie stained chicken liver scanned at 10µm with the 488nm, 520nm and 658nm lasers of the Sapphire Biomolecular Imager.

Pelargonium Stem. Two channel composite image of a Pelargonium Stem scanned at 10µm with the 658nm and 785nm lasers of the Sapphire Biomolecular Imager.

Pine Cone. Multichannel scanned image of a pine cone sliver, scanned with 488nm and 785nm lasers on the Sapphire Biomolecular Imager using the demo machine in Germany.

Cow Stomach.

Chicken Heart.

29

30

3 WESTERN BLOTTING TIPS AND GUIDELINES
31

WESTERN BLOTTING TIPS AND GUIDELINES

3.1. Tips for Successful Western Blot Transfer
Whether you choose to do a wet or dry transfer, there are many steps you can take to maximize transfer efficiency.

Wet Transfer
Benefits of Wet Transfer
Using a wet transfer apparatus, high intensity (1-2 hour) or lower intensity (overnight) transfers can be performed. This allows for optimizing transfer conditions for each individual protein. Transfer times can be shortened to prevent transfer of low molecular weight proteins through the membrane while longer transfer times can be used to promote complete migration of high molecular weight proteins out of the gel.

Transfer “stack”

Cassette

Tank

Drawbacks to Wet Transfer

The transfer process generates heat, which can decrease the resistance of the transfer buffer resulting in inconsistent transfer across the gel. High heat can also result in breakdown of the gel itself. To prevent heating, transfer buffer should be pre-chilled prior to use. In addition, the transfer buffer should be kept cold during transfer. Long transfers are often performed in a cold room to aid in keeping the buffer cool. Additionally, selfcontained ice blocks can be placed within the tank and are usually supplied with the apparatus. Alternatively, external-cooling mechanisms can be used to control heating.

Advantages
· Flexibility: · Transfer conditions are easily adjusted · Multiple transfer buffer options enable more ways to optimize transfer
· Supports transfer of a broad molecular weight range at one time
· Compatible with extended transfer times for large molecular-weight proteins
· Can be used for quantitative Westerns

Disadvantages
· Heating of buffer can interfere with transfer · Cooling mechanism and/or cold room space required
during transfer · Large volumes of transfer buffer are required

Semi-Dry Transfer

Benefits of Semi-Dry Transfer

Semi-dry transfers are fast and easy and

require little buffer. Transfer of more difficult to transfer proteins can be optimized by using a

Electrode Plate

+

discontinuous buffer system, a feature unique

to semi-dry transfer systems. In a discontinuous

buffer system, the filter paper placed on the

Electrode Plate

­

anode side of the stack is soaked in a different

buffer from the filter paper placed on the

cathode side of the stack. This can increase

migration of the protein out of the gel while also

promoting better retention on the membrane.

+
­ Power Cord

Apparatus Cover Transfer “stack”
Apparatus Bottom

32

WESTERN BLOTTING TIPS AND GUIDELINES
Drawbacks to Semi-Dry Transfer Semi-dry transfer systems have less flexibility and it can be difficult to transfer both high and low molecular weight proteins. Low molecular weight proteins can transfer through the membrane due to the high intensity blotting conditions while high molecular weight proteins may not efficiently transfer out of the gel due to decreased transfer times. However, transfer of high molecular weigh proteins can be improved by using the discontinuous buffer system. Transfer times cannot be extended when using semi-dry transfer, as there is limited buffering capacity. In addition, the gel can dry out if insufficient buffer is used.

Advantages

Disadvantages

· Transfer is rapid · Can use discontinuous buffer system to optimize
transfer of proteins · Little buffer is required · Easy to set up · Good for performing large numbers of blots analyzing
the same protein

· High intensity field strength may cause low molecular weight proteins to migrate through membrane
· Difficulty in transferring high (>120 kDa) molecular weight proteins
· Not recommended for quantitative Westerns
· Gel can dry out

Tips for Successful Transfers · Take care when preparing transfer buffers; small inconsistencies can affect transfer · Use high quality, reagent grade methanol when preparing buffers; impurities in methanol can decrease
transfer efficiency · Never dilute the transfer buffer · Do not adjust pH of the transfer buffer · For optimum transfer, do not reuse transfer buffer · Make sure all equipment is clean · Use a thinner gel (0.5-0.75 mm thickness) · If using a PVDF membrane, pre-wet the membrane in 100% methanol prior to equilibration in transfer buffer · Equilibrate transfer pads, filter paper, membrane and gel for at least 15 minutes in buffer(s) · Remove all air bubbles and creases between each layer in the stack · Ensure the stack is firmly held in place and pressure is applied evenly over the entire surface of the stack · Ensure that electrodes are connected and free of debris and damage (plate electrodes) · Make sure the current is applied correctly and proteins will migrate towards the membrane · Use pre-stained molecular weight standards to help monitor transfer · Stain the blot with a reversible total protein stain (e.g. Ponceau S) to check quality of transfer · For difficult to transfer proteins, adjust methanol and SDS concentrations. SDS promotes migration of proteins
out of the gel but can inhibit membrane binding. Methanol increases retention of proteins on the membrane but can hinder migration out of the gel.

33

WESTERN BLOTTING TIPS AND GUIDELINES

Wet Transfer Buffers
Towbin Transfer Buffer · Standard wet transfer buffer · SDS (0.025-0.1%) can be added to facilitate transfer of proteins
25 mM Tris, pH 8.3 192 mM glycine 20% methanol +/- SDS
CAPS (3-[cyclohexylamino]-1 propane sulfonic acid) Buffer · For blotting basic proteins · For blotting prior to N-terminal sequencing
10 mM CAPS, pH 11 10% methanol
Dunn Carbonate Buffer · For higher efficiency transfer of basic proteins · Can be used to enhance ability of antibodies to recognize some antigenic sites
10 mM NaHCO3 3mM Na2CO3, pH 9.9 20% methanol
Semi-Dry Transfer Buffers
Bjerrum Schafer-Nielsen Buffers · Standard semi-dry transfer buffer based on Towbin
48 mM Tris, pH 9.2 39 mM glycine 20% methanol
Discontinuous Tris-CAPS Buffer System · Uses two different buffers to enhance transfer of proteins · The filter paper assembled on the membrane side (anode) of the blot contains methanol · The filter paper on the gel side (cathode) of the blot contains SDS
60 mM Tris, pH 9.6 40 mM CAPS + either 15% methanol or 0.1% SDS

3.2. How to Improve Your Chemiluminescent Western Blots
Chemiluminescent Westerns are popular assays for assessing protein expression. In this indirect detection method, chemiluminescent substrates emit light when reacted with an antibody conjugated to an enzyme. The emitted light is captured and archived on x-ray film (traditional), or through digital imaging. Chemiluminescent Western blotting is a highly sensitive assay and can detect femtograms of protein. Although it is only semiquantitative, it is useful for detecting the presence or absence of

Substrate
HRP Secondary Antibody

Protein Sample

Primary Antibody

34

WESTERN BLOTTING TIPS AND GUIDELINES
a protein. For example, chemiluminescence can be used to detect the induction of exogenous protein expression, to confirm and follow purification of a known protein, or for verification of antibodies during production. Chemiluminescent Westerns can be difficult to perform. The ultimate goal is to obtain a blot with a high signal-tonoise ratio. However, chemiluminescent Westerns can be plagued with high background, either in the form of an overall background that masks the signal from the protein of interest, or as bright dots and speckles and/or splotches scattered randomly over the blot. The increase in background noise can arise from a variety of factors. It can also be difficult to obtain a strong signal from the protein of interest. Tips for Improving Chemiluminescent Westerns · Optimize the amount of protein to load on the gel. In general, 20-40 g of total protein can be loaded without
overloading the well. However, the total amount of protein should be optimized for each protein:antibody pair. Ideally, enough protein should be loaded to allow for easy capture of the signal without experiencing saturation. · Choose the correct membrane. Nitrocellulose and PVDF membranes are commonly used, and each has advantages and disadvantages. · Keep everything clean. Prevent background by thoroughly cleaning all equipment and trays prior to use. Only handle the gel and membrane with gloved hands. Keep trays covered during incubations. · Ensure that all buffers are well mixed. Particulates in blocking and antibody incubation buffers will stick to the membrane and cause high background. Buffers can be filtered prior to use. · If experiencing high background, use a larger volume of washing buffer and increase number and duration of washes. · Try different blocking buffers. Some antibodies react with proteins in blocking buffers, causing a high background. Alternatively, some blocking buffers can mask the protein of interest, preventing detection. Nonfat dry milk and bovine serum albumin are the two most common protein blockers containing multiple proteins. Blocking buffers containing one protein can also be used. Protein-free buffers can be used when the primary antibody reacts with protein components in the buffer. · Titrate both primary and secondary antibodies. Use a dot blot and checkerboard titration to determine the optimum primary and secondary antibody concentrations. · Never dilute a horseradish peroxidase-conjugated secondary antibody in buffer with sodium azide. Sodium azide inhibits HRP activity. · Use enough substrate. Make sure the blot is coated entirely with substrate to prevent local concentration differences. · Try different substrates to increase sensitivity and signal duration. Different substrates are available with differing sensitivities for detecting high to moderate versus low abundance proteins. Different substrates also have different reaction rates and emit light for different durations of time. This can affect your ability to capture multiple exposures. · The substrate may need to be equilibrated to room temperature before use to increase the enzyme activity. · Use a digital imager rather than film. Digital imagers increase the linear dynamic range allowing easier detection of low abundance proteins while limiting saturation when detecting high abundance proteins.
35

WESTERN BLOTTING TIPS AND GUIDELINES

3.3. How to Improve Your Fluorescent Western Blots

Fluorescent Western blots are the gold standard for quantitative Westerns. They are ideal for detecting multiple proteins simultaneously (multi-plexing), allowing in-lane normalization and detection of your protein of interest and loading control at the same time. In addition, post-translational modifications can be studied and quantitated easily.

Protein A

Secondary Antibody
Primary Antibody

Secondary Antibody

Protein B

Primary Antibody

In fluorescent Western blotting, the secondary antibody is directly conjugated to a dye, which is excited by light. The emitted light is detected by a digital imager and digitized for data analysis. Multiple proteins can be detected simultaneously by using secondary antibodies conjugated to different dyes with non-overlapping spectral emissions.

Although similar to chemiluminescent Westerns, there are additional factors that must be taken into consideration when performing fluorescent Westerns:
· Titrate both primary and secondary antibodies. Use a dot blot and checkerboard titration to determine the optimum primary and secondary antibody concentrations.
· When multi-plexing, optimize detection of each target separately prior to simultaneous detection.
· Primary antibodies may need to be increased 2-5x compared to concentrations used in chemiluminescent Westerns.
· Secondary antibodies may also need to be increased (1:5000 is a recommended starting dilution).
· Use a PVDF membrane with low autofluorescence. Nitrocellulose and some PVDF membranes can autofluoresce causing high background.
· Avoid inks and dyes that can fluoresce. Use a pencil to mark the blot. Common dyes such as bromophenol blue and Coomassie autofluoresce.
· Keep everything clean. Prevent background by thoroughly cleaning all equipment and trays prior to use. Only handle the gel and membrane with gloved hands. Use powder-free gloves. Keep trays covered during incubations.
· If you are using fluorescent molecular weight markers, skip a lane before loading samples. This will prevent the signal from the molecular weight markers from bleeding into sample lanes.
· Work with fluorescent antibodies on the bench top, but store stocks in the dark.
· When multiplexing, use primary antibodies made in different species and secondary antibodies that are highly cross-adsorbed to prevent cross recognition.
· Avoid spectral overlap when multiplexing. Choose fluorophores that have optically distinct spectra.
· To increase the specific signal, always detect the strongest target in the blue channel, the middle in the green channel and the weakest in the red channel.
· When archiving blots, store them in the dark.

36

WESTERN BLOTTING TIPS AND GUIDELINES

3.4. Transition from Chemiluminescent to Fluorescent Western Blots
Bring Fluorescent Western Blotting to Your Lab. It’s Easier Than You Think.
Chemiluminescence is the most familiar method of detection for Western blotting and offers great sensitivity. However, many scientific questions and experimental designs require the additional information provided by fluorescent Western blotting; this includes precise quantitation and visualization of similarly sized proteins within the same sample.
Once you make the decision to move to fluorescent Western blotting, what comes next? First, you can check if fluorescent Western blotting is right for your experiment using the flow chart in the figure below. Then, refer to the tips and advice provided in this document to get started with your first fluorescent Western blot.

Do you have access to a digital imager?

No

Yes

Do you have multiple proteins
to detect on the same blot?

No

Yes

Are they similar sizes?

No

Yes

Do you anticipate a very low level
of protein?

No

Yes

Do you need precise quantitation?

No

Yes

Chemiluminescence

Either

Chemiluminescent or Fluorescent Western Blotting.

Fluorescence

Before You Run the Gel
Dilute Ladder
Some pre-stained protein ladders fluoresce strongly and can interfere with detection of proteins expressed at a low level. Try diluting the ladder 1 to 10 before loading.
Optimize Sample Concentration
As with any new assay, quality results are dependent on first optimizing your assay for the samples to be used. Sample load volume may need to be adjusted for the best possible results. When first switching from chemiluminescent to fluorescent Westerns, try a slightly higher sample load volume to confirm that your assay is working. Once you have a positive result, you can lower the sample load volume and push for more sensitive results.

Make considerations for the protein ladder, sample concentrations, and loading dye.

37

WESTERN BLOTTING TIPS AND GUIDELINES
While Running the Gel Be Wary of Loading Dye Many loading dyes can fluoresce. To prevent this from interfering with the signal on a fluorescent blot, run the dye front off the gel or cut it off before transfer.

1 2 3 4 5 6 7 8 9 10

Transferring to the Membrane PVDF Membranes For the best signal-to-noise ratio, use a membrane with minimal autofluorescence such as Azure’s low fluorescence PVDF membranes. Membrane Handling Any contamination to the membrane will be obvious in a fluorescent Western blot. Always use forceps when handling membranes. Products · Pre Cut PVDF (AC2108 ­ AC2109)

Before setting up the transfer, make sure to trim off the loading dye if possible to prevent it from interfering with your data.
Use PVDF membranes and handle only with forceps.

Probing the Membrane
Blocking
As with chemiluminescent blots, it is important to completely block the membrane to avoid nonspecific binding of antibodies and to reduce background. Azure Protein Free Blocking Buffer and Azure Fluorescent Blot Blocking Buffer are formulated to not only reduce background, but also to stabilize the fluorescent dyes of AzureSpectra secondary antibodies for enhanced signal detection.
Optimize Antibodies
Antibody concentration may need to be optimized and adjusted for the best possible results. When first switching from chemiluminescent to fluorescent Westerns, use the manufacturer’s recommendations for antibody dilutions.
Incubation Trays
Fluorescent secondaries can become quenched when exposed to bright light for long periods of time. When incubating your sample with light sensitive antibodies, cover your blot with an opaque material (such as with Azure’s opaque incubation trays) to protect from quenching.
Products
· Azure Protein Free Blot Blocking Buffer (AC2112) · Azure Fluorescent Blot Blocking Buffer (AC2190) · Fluorescent secondary antibodies (AC2128 ­ AC2139,
AC2156 ­ AC2171) · Opaque Incubation Trays (AC2120 ­ AC2123)

Detector Molecule

Secondary Antibody

Primary Antibody
Target Membrane
Diagram of the antigen-antibody and antibody-antibody interactions in Western blotting detection.

Tip: use opaque containers to protect the light-sensitive fluorophores during incubation.

38

WESTERN BLOTTING TIPS AND GUIDELINES

Washing
Stringent Wash
Because the fluorescent dyes can adhere to the membrane, washing is extremely important to reduce background. Make sure to use a stringent wash, especially when working with near-infrared fluorophores. Azure Fluorescent Blot Wash Buffer is specially formulated for use with fluorescent Western blots.
Wash Volume
The volume and duration of washing is also important to rid the membrane of any free dye and antibody. We recommend two quick rinses in 25mL wash buffer followed by three 5 minute washes in 25mL wash buffer.
Final Rinse
Any detergent in the wash buffer can also fluoresce, so rinsing in PBS or TBS after the final washing step is essential to lower background signal.
Products
· Azure Blot Washing Buffer (AC2113) · Azure Fluorescent Blot Washing Buffer (AC2145)

To prevent background signals and get the clearest data, follow the recommended steps such as thorough washes after antibody incubation.

Imaging Reduce Background Contamination When many people use the same imager, the insides and trays can become contaminated. Background Quenching Sheets absorb background fluorescence to improve signal-to-noise ratios. Products · Quenching sheets (AC2144, AC2147)

To prevent background fluorescence from interfering with your signal, use Azure’s quenching sheets.

Want some help getting started?
A demo kit is a great way to try out fluorescent Western blotting in your own lab with your own samples and primary antibodies. It contains everything you need to perform single color fluorescent Western blotting:
· PVDF Membrane · Fluorescent Blot Wash Buffer · Fluorescent Blot Blocking Buffer · Secondary Antibody · 2 Quenching Sheets
AzureSpectra Demo Kits · Goat -Mouse 700 (AC2172) · Goat -Rabbit 700 (AC2173) · Goat -Mouse 800 (AC2174) · Goat -Rabbit 800 (AC2175) · Goat -Mouse 650 (AC2176) · Goat -Rabbit 650 (AC2177) · Goat -Mouse 550 (AC2178) · Goat -Rabbit 550 (AC2179)

39

WESTERN BLOTTING TIPS AND GUIDELINES

3.5. Western Blot Normalization
Quantitative Westerns: What is the Best Way to Normalize Your Western blot?
Far from being an “is-it-there-or-not” technique, modern digital detection instruments can make Western blotting reproducible and quantitative. By working within the linear dynamic range of your detection method and normalizing the data to control for variations in protein load and membrane transfer, you can get truly quantitative results. But what is the best way to normalize a Western blot? In the past, the gold standard normalization method was to use a housekeeping protein based on the assumption that the levels of these proteins are fairly consistent across experimental conditions and cell lines. However more recent studies have shown that this assumption is not always true1,2 leading to inaccurate measurements of relative protein abundance. Instead, quantitative Western blotting experts1,2 and the journals they publish in4 are recommending a new gold standard for normalization–normalizing to total protein detected in each lane, preferably by staining on the membrane.
Western Blot Normalization: Housekeeping Protein vs Total Protein

Normalization method Housekeeping protein

Total protein

Benefits

· Familiar, commonly used

· Large linear dynamic range · Low variability · Consistant across sample types · No change with experimental conditions

Challenges

· Narrow linear dynamic range
· Abundance can vary with experimental conditions
· Abundance may not be consistent between sample types
· High variability
· Must ensure housekeeping protein physically resolved from protein of interest on gel

· Must ensure total protein stain used is compatible with antibody binding and detection method

Using Housekeeping Proteins for Normalization
The most significant drawback of using housekeeping proteins is that their levels may not be consistent across samples and conditions.1,2 It is possible to use a housekeeping protein for normalization, but you must first spend the time and effort to validate your choice, and may need to examine multiple potential standards before you find one that is truly expressed at the same level across all of your samples and does not change across your experimental conditions.
A second significant challenge associated with housekeeping proteins is their high abundance.1,3 If the housekeeping protein is present at a very high level in your sample, this limits the amount of sample you can load on the gel because you will need to keep the housekeeping protein within the linear range of detection and not saturate the signal for the housekeeping protein. This is particularly problematic if the protein of interest is not similarly highly expressed because the two proteins will not be within the same linear range of detection.2,3
A third challenge to consider if you’re doing multiplex Western blots–such as comparing phosphorylated and non-phosphorylated forms of the same protein–is the complexity of generating primary and secondary antibodies from non-overlapping species.
Finally, it is always possible that detecting the housekeeping protein could interfere with detection of the protein of interest.1 Ideally, the housekeeping protein should be a different size than the protein of interest so the two proteins are spatially resolved on the blot. This becomes increasingly difficult when an experiment examines multiple proteins of interest on the same blot.

40

WESTERN BLOTTING TIPS AND GUIDELINES

Using Total Protein Staining for Normalization

With total protein normalization, instead of trying to find a protein that can represent the total amount of sample that transferred to the membrane, total protein is measured on the membrane directly and this value is used as the denominator when normalizing.1-4 Many total protein stains are available that can be used to stain gels and membranes.1 Total protein stains provide a larger dynamic range and demonstrate lower variability and cleaner data than housekeeping proteins.1,2

Traditional Western Blot Workflow
Separate proteins on gel
Transfer proteins to membrane

Western Blot Workflow with AzureRed TPS Separate proteins on gel
Transfer protiens to membrane
Stain for total protein with AzureRed (30-minute protocol)

Total protein normalization can be much faster than using a

Block membrane

Block membrane

housekeeping protein, especially for chemiluminescent blots because the staining step takes less time that stripping and

Finish Western blot protocol

Finish Western blot protocol

reprobing the blot. Ideally, total protein staining is conducted

on the membrane, either before or after immunodetection.2

With some stains such as AzureRed Fluorescent Total Protein Stain, it is possible to stain the blot before

immunodetection and then to image total protein simultaneously with the protein(s) of interest. With this simplest of

workflows, images for the protein(s) of interest and total protein are automatically aligned, avoiding the need resize

and align images captured at different times.

The analysis workflow after image capture is essentially unchanged compared to using a housekeeping protein; the signal density for the entire lane or a large portion of the lane is used for normalization instead of the density for a single band.

Staining the membrane with a total protein stain provides an added quality control benefit, allowing verification that membrane transfer was complete and free of artifacts.

Looking for More Information About Western Blot Normalization?
Please see the Western Blot Normalization application note to learn more about the different methods of Western blot normalization and to help decide which normalization approach is best for your blot. Read more about using AzureRed Fluorescent Total Protein Stain for total protein normalization of fluorescent Western blots in the Accurate Western Blot Normalization With AzureRed Fluorescent Protein Stain application note.

References
1. Moritz CP. Tubulin or not tubulin: heading toward total protein staining as loading control in Western blots. Proteomics. 2017;17:1600189.
2. Thacker JS et al. Total protein or high-abundance protein: which offers the best loading control for Western blotting? Anal Biochem. 2016;496:76-78.
3. McDonough AA et al. Considerations when quantitating protein abundance by immunoblot. Am J Cell Physiol. 2015;308(6):C426-C433.
4. Fosang AJ, Colbran RJ. Transparency is the key to quality. J Biol Chem. 2015;209(50):29692-29694.

41

WESTERN BLOTTING TIPS AND GUIDELINES

Images Used: 2015-1228-164847

Images Used:

2016-0105-153125

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The Problem:
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Wash Prior to Stripping

Insufficient or

· Rinse blots with ultra

Wash Prior to Stripping incorrect wash prior to stripping

pure water for at least 5 minutes before stripping

· Do not wash with wash

buffer

·

Ensure adequate buffer and stripping time

volumImeages Used:
2015-1222-1630

2015-1230-1637

2016-0105-1531

The Fix:

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The Problem: The Problem: Not washed before stripping

· Rinse blots with ultra

Washed with wash

· RpuinrseewbalpmotuetinsrreuwfotwiertashatbteuelrletffI2roamo0ars1aret5g-ae51sst2tU0rls9eie-p0da9:ps1ti1n15g2

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Marker too bright

· Cover the marker before imaging

· Allow blot to dry before

Images Used: 2015-1209-091 2016-0107-101
The ·FRixin:se blots with ultra
pure water for at least 5 minutes before stripping

imaging

· Do not wash with wash

· Cover the marker befo·reDilute marker before Not wimasahgeidngbefore stripping lWoaasdhinegd with wash buffer before stripping

buffer

Images Used:

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Bubbles in Transfer

· Allow blot to dry before

imaging

Images Used:

The Problem: ·

Dilute

marker

before

loading

2016-0104-161425 2016-0107-095429

2016-0107-101420

Marker Too Bright

BubbTlehs ein tFraixns:fer · Press and smooth filter

· Cover the marker befo22r00e1165–01120079–100911 imaging
·TAhlleowFbixlo:t to dry before
imaging

paper after covering

Tips for transferring proteins from gel to membrane to remove

· Dilute marker before loading

membrane

bubbles · Ensure adequate transfer

1. Optimize voltage and duration of transfer for each device

buffer is present in

2. Remove air bubbles before transfer

cartridge

3. Ensure transfer stack has a tight fit

· Press and smooth filter paper

after covering membrane to

Transfer stack was not tight: Replace sponge pads

Air bubbles: Ensure bubbles are removed or degas transfer buffer

Contamination: Clean removFeinbguerbpbrlientss: Always use equipment and replac·e Ensure adfoerqceupastteothraanndslefer
sponge pads buffer is premseenmtbirnanceartridge

· Cover the marker before imaging
· Allow blot to dry before

imaging

· Dilute marker before loading

42

WESTERN BLOTTING TIPS AND GUIDELINES

The Problem:
InsufficienItmoargeUneven Transfer Pressure

Images Used: 2016-0104-164834 2016-0111-171150
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· Place gel and membrane

in the middle of the casette · Transfer set up should be
very tight

· Place gel and membrane in

the middle

Images Used: 2015-1230-164415

2016-0106-162523

The Problem:

The Problem:

2015-1214-151723

Antibody Cross-reactvity AntiTbhoedFyixC: ross-reactvity

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· Avoid antibodies from

cross-reactvity

species that are too “close”

(ex. rat and mouse)

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Resulting images
The Fix:

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· Block in between antibodies t2h0a1t5-1216-151617 might cross react

antibody

· Other causes:

· Non-specific binding

· Too much antibody

The Problem:
High Background

· Insufficient blocking

Images Used:

2016-0106-161352 2016-0107-094635

Images Used: 2016-0106-161352 2016-0107-094635

· ScreTehneaPnrtoibbloemdi:es

· OtThheHreiCgFahiuxBs:easc:kNgoronu-Snpdecific T· hSecrFeiexn: antibodies

Binding High background · Use low-fluorescence
· Too much·anmUsteeimbcboorrdarenycet washing

· Other Causes: Non-Specific Binding

· Insufficient pbrlootocckoilnagnd wash buffer
· After washing rinse with
Tips for preparing samples and loadingwagteerls

· Too much antibody · Insufficient blocking

· Do not use excessive

1. Ensure samples are free of cellular debris and are not viscous

amounts of antibody

2. Load samples slowly and make sure not to overload the wells 3. Optimize the amount of protein loaded

· Dry the membrane before

· The usual: memimbargainnge, blocking,

Cellular debris in samples: Sonicate and filter

syDriNngAeiwnif sinaamssaphmlepisnl:eSgbhue,fafrreisrnaomrspaidnledsgDw,Nitaahsentibody load

· The usual: membrane, blocking, washing, rinsing, antibody load
· Dry the membrane before imaging

Cellula·r dDerbyritshien mem·bSroanniecabteefaonrdefimltearging

250 Cell SV1 SV2 SV3 S1 S2 S3 S4 S5

samples

· For DNA in samples, shear

150

samples with syringe or

100 75

add DNTarasnesferrin

50

Actin

37

GAPDH

25 20

15 10

43

44

4 PROTOCOLS
45

PROTOCOLS

4.1. HRP/Chemiluminescent Blot Detection with Radiance Substrate
Long Protocol for Catalog Numbers
· AC2100–Radiance Sample, sufficient for 140 cm2 membrane · AC2101–Radiance, 150 ml, sufficient for 1500 cm2 membrane
Additional Materials Required · Electrophoresis apparatus and buffers for SDS-PAGE · Tank and buffers for electrophoretic transfer of proteins from gel to membrane · Nitrocellulose or PVDF membrane, cut to size of gel. All membrane products available from Azure (see the
Related Products section) are compatible with Radiance. · Washing buffer (PBS-T or TBS-T). For best results, use Azure Fluorescent Blot Washing Buffer (see the Related
Products section) · Blocking buffer · Primary antibody compatible with your application · Secondary antibody, conjugated to Horseradish peroxidase (HRP) corresponding to your primary antibody (see
the Related Products section) · CCD-based detection system, or film (see the Related Products section)
Background
Radiance chemiluminescent horseradish peroxidase (HRP) substrate is specially formulated for CCD imaging and to provide excellent performance with Azure Imaging Systems. Radiance produces a strong, long-lasting signal, which, combined with very low background levels, allows for long exposure times enabling the detection of lowabundance proteins. Additionally, the signal from Radiance is linear with respect to protein amount over a broad range of concentrations, displaying no substrate depletion at high protein loads, allowing the user to take full advantage of the linear range of the CCD detection method. Radiance is also compatible with X-ray film detection, though the limited dynamic range of film will make resulting data less quantitative.
Since 1988, enhanced chemiluminescence or ECL (1) has become one of the most common detection methods in Western blotting (2). In this method, the secondary antibody is conjugated to the enzyme Horseradish peroxidase (1,2). Once bound to the membrane, the secondary antibody is detected by incubating the blot with a solution containing an HRP substrate that generates a light-emitting product after reaction with HRP (Figures 4.1, 4.2). The chemiluminescent signal can be detected by exposing the blot to X-ray film, or by imaging with a CCD camera. Radiance is an enhanced chemiluminescent substrate specially developed for CCD imaging. Radiance produces a bright signal with very low background for extremely high sensitivity and a detection limit of attomoles of protein. Additionally, the Radiance signal is long lasting, which combined with the low background, allows long-term exposures to detect low-abundance proteins.

Substrate
HRP

Light

Oxidized Products

Secondary Antibody HRP Conjugate

Primary Antibody

Membrane

Proteins transferred to the membrane

Blocker

NH

O

NH HRP + HO
NH

O
Luminol

NH

O

O+ N + light
O-
O

Figure 4.1. The principle of chemiluminescent Western blotting.

Figure 4.2. Chemiluminescence of luminol.

46

Overview of the Protocol for Chemiluminescent Western Blots
Electrophoresis to separate proteins in sample
Transfer proteins from gel to membrane
Block to mask nonspecific protein binding sites on membrane
Primary antibody binds to protein of interest
Wash to remove excess antibody
Secondary antibody binds to primary antibody
Wash to remove excess antibody
Substrate (Radiance) substrate reacts with HRP bound to secondary antibody
to create luminescent signal
Image detect luminescent signal with CCD camera or film

Quick Protocol For additional information, see the detailed protocol which follows.
Step 1. Prepare your protein blot
2. Block membrane for 1 hour at room temperature (RT)
3. Incubate blot with primary antibody for one hour at RT with gentle agitation
4. Wash blot: · 1 x quickly · 1 x 15 min, with 1 ml/cm2 membrane · 3 x 5 min, with at least 0.5 ml/cm2 membrane each time
5. Incubate blot with secondary antibody for one hour at RT with gentle agitation
6. Wash blot: · 3 x 5 min, with at least 0.5 ml/cm2 membrane each time
7. Mix Radiance components 1:1 and place 0.1 ml/cm2 on blot for 2 minutes
8. Drain excess reagent
9. Cover damp blot with plastic wrap and image by CCD camera or exposure to X-ray film

User Notes

PROTOCOLS
47

PROTOCOLS

Detailed Protocol

Step

Notes

1. Prepare a protein blot

1.1. Separate the protein sample(s) via electrophoresis
1.2. Transfer proteins to membrane
· Prewet membrane in transfer buffer, and assemble transfer sandwich according to tank manufacturer’s instructions.
· Dot-blots or slot blots can also be detected with Radiance.

· Any electrophoresis system and buffer, such as Laemmli, is compatible with Radiance.
· A wet or tank transfer method is preferred, though semi-dry methods should also be compatible. We have found that the buffer system developed by Bolt et all (3) works well.
· Both nitrocellulose and PVDF membranes are compatible with Radiance.
· If using PVDF, first wet membrane with a 1 min incubation in 100% MeOH followed by water for ~5 min and then transfer buffer for 5-10 min.
· For slot blot applications, nitrocellulose is much more convenient than PVDF because it is more difficult to avoid bubbles with PVDF.

2. Block membrane

· Incubate the blot in a blocking buffer with gentle agitation for 1 hour at room temperature (RT). Use 0.2 to 0.5 ml of blocking buffer per cm2 of blot to provide adequate blocking.

· Blocking masks non-specific protein binding sites on the membrane, reducing background and increasing the specificity of binding of the primary antibody to the protein of interest.
· The optimal blocking buffer will depend in part on the nature of the antigen of interest, and on the quality of the primary antibody. Common blocking agents including nonfat dry milk have been found to be compatible with Radiance.
· 10 to 20 ml is usually sufficient for a typical 7 x 9 cm mini-blot.

3. Incubate blot with primary antibody

· Dilute primary antibody in blocking buffer.
· Incubate blot with primary antibody solution for 1 hour at RT with gentle agitation.

· Optimal primary antibody dilutions must be determined empirically.
· For CCD imaging, we recommend primary antibody dilutions from 1:1000 to 1:10,000. A good initial dilution is 1:5000.
· If the blot will be imaged on film, use 2­5x less primary antibody than for CCD imaging. For example, if 1:1000 dilution of the primary antibody was optimal for CCD detection, 1:5000 is suitable for film detection.
· Antibody can be added to a dish and placed on a shaker, or a smaller volume (5-10 ml) can be used by sealing the blot into a bag and placing it on a rotary platform.

4. Wash blot to remove excess primary antibody

· 1 x quickly
· 1 x 15 min, with 0.7 ml/cm membrane
· 3 x 5 min, with at least 0.3 ml/cm2 membrane each time.

· For best results, use Azure Fluorescent Blot Washing Buffer (AC2113) which is optimized for chemiluminescent as well as fluorescent blots. PBS-T or TBS-T are also compatible with Radiance.
· We recommend washing or rocking blots in a clean dish on a shaker to provide gentle agitation.
· For example, a standard 7×9 membrane requires: ~50 ml of washing solution for the 15 min wash; and ~20 ml of washing solution for 5 min washes.

5. Incubate blot with secondary antibody

· Dilute secondary antibody in blocking buffer.
· Incubate blot with secondary antibody solution for 1 hour at RT with gentle agitation.

· Optimal secondary antibody dilutions must be determined empirically.
· We recommend secondary antibody dilutions of 1:5,000 to 1:20,000. A good initial dilution is 1:10,000.
· If the blot will be imaged on film, use 2­5x less secondary antibody than for CCD imaging. 1:50,000 dilution is a good starting point for film detection.
· See also notes for step 3.

6. Wash blot to remove excess secondary antibody

· 3 x 5 min, with at least 0.3 ml/cm2 membrane each time.

· See notes for step 4.

48

PROTOCOLS

7. Incubate blot with Radiance

· Mix components 1 and 2 in a 1:1 ratio in sufficient amounts to obtain at least 0.1 ml/cm2 of the blot and add to the blot.
· It is better to prepare the working solution just before use. However, mixed reagent is stable for several hours at RT.
· Allow substrate to react with blot for 2 minutes.

· Be careful not to touch or put pressure on the blot as this can result in non-specific background.
· Use only plastic forceps, not metal; metal forceps damage the blocked surface, creating new adsorption sites. Also, traces of metal may act as a catalyst for nonenzymatic substrate oxidation, resulting in very high background.
· The minimal amount of working reagent is 0.1 ml/cm2. For example, for a 7 x 9 cm blot, this minimal volume is 7x9x0.1=6.3 ml.
· If using the minimal amount of working reagent, incubation may be done without agitation. Make sure the membrane surface is level so adequate reagent is held by surface tension.
· Incubation may also be done with gentle agitation in a tray just slightly larger than the membrane. Increase the reagent volume as necessary to ensure the membrane is adequately covered with reagent.

8. Drain excess reagent

· Remove excess substrate via capillary action by touching a KimWipe® or other absorbent material to the edge of the blot.

9. Image blot

· While blot is damp, cover with transparent plastic wrap and either place blot in CCD imager, or expose blot to film.

· We recommend trying three exposures; 30 sec, 2 min, and 5 min.
· The blot can be imaged and re-imaged for several hours; 70% of the initial signal will remain after 60 minutes, and substantial signal will remain after 8-10 hours.

Troubleshooting & FAQ
Western blotting can require substantial optimization due to the multiple steps involved. The correct amount of protein to load on the gel and the best dilutions of primary and secondary antibodies must be determined empirically. Some common questions are addressed below:

Problem High background
No or low signal
White spots within bands Speckled background

Possible Soutions
· Reduce primary antibody concentration by increasing the dilution factor. · Try a different blocking buffer. · Try a shorter exposure time. · Increase washing time.
· Check that correct primary antibody used. · Check that secondary antibody recognizes primary (for example if the primary is a rabbit
antibody, that the secondary is goat-anti-rabbit).
· Improve transfer, making sure to remove any bubbles between the gel and the membrane.
· Filter secondary antibody. · Filter blocking and washing buffers. · Ensure that the laboratory environment is clean, to minimize dust, debris or any other
particles that might come in contact with the blot. Cover the dish during incubation or washing steps. · Use non-powdered gloves, or switch to a different kind of gloves. We recommend powderfree nitrile gloves or polyethylene gloves.

49

PROTOCOLS

References
1. Thorpe GH, Kricka LJ, Moseley SB, Whitehead TP, Phenols as enhancers of the chemiluminescent horseradish peroxidaseluminolhydrogen peroxide reaction: application in luminescencemonitored enzyme immunoassays. Clin Chem. 1985 Aug; 31(8): 1335-41.
2. Leong MM, Fox GR., Enhancement of luminol-based immunodot and Western blotting assays by iodophenol. Anal Biochem. 1988 Jul; 172(1): 145-50.
3. Bolt M.W., Mahoney P.A, High-efficiency blotting of proteins of diverse sizes following sodium dodecyl sulfate-polyacrylamide gel electophoresis. Anal Biochem. 1997 May 1; 247(2): 185-192.
Related Products

Catalog Number AC2113 AC2114 AC2115 AC2105 AC2106 AC2107

Product Azure Fluorescent Blot Washing Buffer Goat-anti-rabbit HRP-conjugated secondary antibody Goat-anti-mouse HRP-conjugated secondary antibody Low Fluorescence Western Membrane (PVDF) 7×9 cm Nitrocellulose Transfer Membrane 0.45 m 7×9 cm Nitrocellulose Transfer Membrane 0.22 m 7×9 cm

Size 500 ml 500 l 500 l 10 sheets 10 sheets 10 sheets

50

PROTOCOLS
4.2. Fluorescent Western Blot Protocol with AzureSpectra Reagents
Quantitative, multi-color and near-IR fluorescent Western blotting kits. Long Protocol for Catalog Numbers · AC2193–AzureSpectra rb650/ms550 Kit · AC2191–AzureSpectra IR rb700/ms800 Kit · AC2192–AzureSpectra IR ms700/rb800 Kit
All kits · Low-fluorescence PVDF transfer membrane 9×7 cm. 10 each · Background Quenching Sheets, 10 each · Azure Fluorescent Blot Blocking Buffer, 1x ready-to-use solution, 300 ml · Azure Fluorescent Blot Washing Buffer, 10x concentrate, 250 ml For AC2193 · Goat-anti-rabbit IgG 650, 40 l · Goat-anti-mouse IgG 550, 40 l For AC2191 · Goat-anti-rabbit IgG IR700, 40 l · Goat-anti-mouse IgG IR800, 40 l For AC2192 · Goat-anti-mouse IgG IR700, 40 l · Goat-anti-rabbit IgG IR800, 40 l
Shipping and Storage Conditions Before opening, the kit may be stored at +4°C. AzureSpectra fluorescent secondary antibody conjugates may be stored at +4°C for up to a month. For longer term storage, store antibody conjugates at -20°C. Gently mix and briefly spin the tubes before taking aliquots. Azure Fluorescent Blot Blocking Buffer must be stored at +4°C. Azure Fluorescent Blot Washing Solution can be stored between 4°C and 25°C. Do not dilute excessive amounts of the concentrates to the final working concentration. Prepare only as much as you need for each assay. Store PVDF membranes at ambient temperature in a sealed bag protected from light and moisture. Store background quenching sheets in a sealed bag at ambient temperature.
Additional Materials Required · Primary antibodies: mouse and/or rabbit IgG. · Electrophoresis apparatus, power supply, gels and buffers for standard Laemmli SDS-PAGE. · Electro-blotting apparatus and transfer buffer (see Reference 4). · Methanol. · 1x PBS or TBS without Tween. · Forceps with flat smooth tips. Plastic (non-metal) forceps are strongly recommended. · Powder-free gloves compatible with fluorescent applications. Polyethylene gloves are strongly recommended. · Incubation and washing trays with smooth interiors such as those provided by Azure (product numbers AC2120,
AC2123, AC2150 and AC2153). · Rotary or rocking platform ­ a rocking platform is preferred as mixing with a rocking action typically generates
more uniform background than orbital shaking. · AzureSpectra rb650/ms550 Kit (AC2193): Fluorescent imager compatible with dyes excitable in green
(530nm ­ 560nm) and red (600nm ­ 650nm) light. Further details are discussed in the “Imaging” section.
51

PROTOCOLS

· AzureSpectra IR rb700/ms800 and ms700/rb800 Kits (AC2191 and AC2192): an imager compatible with dyes excitable in far-red (630nm ­ 700nm) and near infrared (750nm ­ 780nm) light. Further details are discussed in the “Imaging” section.

Introduction to Multi-Color Fluorescent Western Blotting
Western blotting provides a means to assay the presence and the expression level of a protein of interest in a complex mixture. Proteins are separated electrophoretically, transferred to a membrane substrate, and the protein of interest is detected with specific antibodies(1, 2). The antibody specific to the protein of interest can be directly labeled, for example, using radioactivity, or can be detected by the use of a labeled secondary antibody. Frequently, a secondary antibody conjugated to horseradish peroxidase (HRP) is used, and the location of the protein is detected via chemiluminescence(3).
When using chemiluminescence, only one protein can be detected per blot. Assaying for a second protein requires stripping and re-probing the blot, a time consuming procedure. Imaging a Western blot using fluorescence allows for multiple proteins to be assayed on one blot by using secondary antibodies labeled with fluorophores having unique excitation and emission spectra(3).
The AzureSpectra rb650/ms550 Kit and AzureSpectra IR rb700/ms800 and ms700/rb800 Kits provide the means to assay two proteins on a single Western blot. Each kit contains two secondary antibody conjugates, each pre-labeled with a different fluorescent reporter. Fluorescent reporters utilized in the AzureSpectra rb650/ms550 Kit are excitable in the visible fluorescence range. The excitation and emission spectra of of the AzureSpectra 550 and 650 conjugates are shown in Figures 4.3 and 4.4. Fluorescent reporters used in the AzureSpectra rb700/ms800 and ms700/rb800 Kits are excitable by far-red and near-infrared light. The excitation and emission spectra of the AzureSpectra NIR conjugates are show in Figures 4.5 and 4.6.
Until recently fluorescent reporters were not widely used for Western blotting applications for various reasons. Membranes typically used for Western blotting had high levels of autofluorescence, appropriate imaging instruments were not readily available, and the time required to acquire an image of a typical blot was relatively long. The most important factor that prevented the use of some fluorescent detector molecules in Western blotting applications was that these proteins need to remain hydrated to maintain their high levels of fluorescence. Western blot membranes can dry quickly and lose the water necessary to sustain fluorescence, and, unfortunately, re-hydrating the membrane does not restore the fluorescence of certain detector fluorophores.
Today, all the above problems can be easily addressed. PVDF membranes with low autofluorescence have become available, and there are several choices of fast and high resolution fluorescent imaging instruments including laserbased scanners and LED-based gel documentation imagers. The most difficult problem is to preserve the hydration of fluorescent detector molecules long enough to be able to image the blot within a reasonable period of time.
We have developed the Azure Protein Free Blocking Buffer and Azure Fluorescent Blot Blocking Buffer to address this problem. The blocking solution provided with the kit contains a component that provides an efficient blocking of the PVDF membrane from non-specific protein binding. At the same time,

Absorbence/Intensity

Absorbence/Intensity

Absorbence/Intensity

Absorbence/Intensity

515 nm ex 565 nm em
250 300 350 400 450 500 550 600 650 700
Wavelength (nm)
Figure 4.3. Absorption and emission s

Documents / Resources

azure biosystems Western Blot Ebook [pdf] User Guide
Western Blot Ebook, Western, Blot Ebook, Ebook

References

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