Western Blotting: A Comprehensive Guide

1 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, 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. Features Uses How To Native • P  roteins retain threedimensional structure and folding • Proteins are separated by mass:charge ratio and cross-sectional area of proteins (Stokes radius) • U  se to study protein complexes • U  se when detection antibody recognizes an epitope on the folded protein • O  mit SDS from loading buffer, gel and running buffer • Omit denaturing agent from sample buffer • Do not heat sample prior to loading Denaturing • P  roteins are denatured using SDS (Figure 1.2) and heat • P  roteins are separated primarily by molecular weight • U  se when native conditions are not required • M  ix protein with sample buffer containing SDS and a denaturing agent • H  eat samples prior to loading Table 1.1. Native vs. denaturing gel electrophoresis. 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 10–43 15 12–60 12 20–80 10 30–95 8 50–200 6 Table 1.2. Resolving power of different concentrations of acrylamide. 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. 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.  resolving small proteins in the 1-10 kDa range, a Tris-tricine system is recommended. With a Tris-tricine gel and For 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. 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 • H  eating of buffer can interfere with transfer • C  ooling mechanism and/or cold room space required during transfer • L  arge 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. Apparatus Cover Electrode Plate + + Transfer “stack” Electrode Plate – – Apparatus Bottom Power Cord Figure 1.4. A typical semi-dry transfer setup. 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 • H  igh intensity field strength may cause low molecular weight proteins to migrate through membrane • D  ifficulty in transferring high (>120 kDa) molecular weight proteins • N  ot 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. Component Example Use Buffer Tris, CAPS, Carbonate Support conduction, maintain appropriate pH Alcohol Methanol, ethanol Increase binding of proteins to membrane Detergent SDS Promote migration of proteins out of gel Tris-tricine Up to 9.5 Tricine separates low MW proteins from free SDS Table 1.6. Transfer buffer components. 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. 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. 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 Primary Antibody Target Membrane Target Membrane Indirect Secondary antibody is labeled. Indirect Secondary antibody is labeled. Figure 1.5. Indirect vs. direct detection. 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. 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.  ight-chain Specific—Antibodies that recognize only the light chain portion of the antibody are L 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.  ffinity Purification—Affinity purification uses an affinity column—a column that contains the A 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. 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 Learn how to optimize your chemiluminescent is present relative to another protein? How much of my protein is Western blots on page 35 in our Western in one sample compared to another sample? How do I control for Blot Tips and Guidelines Chapter. sample loading inconsistencies?” We discuss the drawbacks to chemiluminescent detection in the next section. Substrate HRP Secondary Antibody 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 Disadvantages • Sensitive • Easy, familiar chemistry • Compatible with film or digital imaging • S  emi-quantitative • S  ignal dependent on enzyme kinetics  ingle protein only, loading controls require stripping • S and re-probing Table 1.7. Chemiluminescent detection. 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 Find out how to improve your fluorescent based on their specific excitation and emission spectra, enabling Western blots on page 36 in our Western Blot multiplexing (detection of multiple proteins simultaneously, Tips and Guidelines chapter. Figure 1.8) for faster, more efficient studies. Thus, one of 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 Secondary Antibody Primary Antibody Protein A Primary Antibody Protein B 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. 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 Figure 1.9. Multiplex fluorescent Western. 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 c600 (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. Advantages Disadvantages • Multiplex capability • Increased quantitative accuracy • Fluorescent label stability allows blots to be stored and re-imaged later • C  an be less sensitive than chemiluminescence • Membranes auto-fluorescence can increase background Table 1.8. Fluorescent detection. 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. 1.6. 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): 680–685.